To image Rab signaling activity in single dendritic spines, we developed FRET sensors for Rab4, 5, 7, 8, and 10. The Rab sensors have two components: (1) the Rab protein tagged with a fluorescent protein that serves as a FRET donor (monomeric enhanced green fluorescent protein (mEGFP)-Rab or mTurquoise2-Rab) and (2) a Rab binding domain (RBD) from a specific effector protein that is tagged with two fluorescent proteins as FRET acceptors (mCherry-RBD-mCherry or mVenus-RBD-mVenus). When the Rab protein is activated, RBD and Rab increase their binding and thus increase FRET between the donor and acceptor fluorophores (Figure 1a). The fraction of Rab bound to the RBD (binding fraction) was calculated by measuring the fluorescence lifetime of the donor (Yasuda, 2012; Yasuda, 2006).

For Rab4, 5, 7, 8, and 10, we used Rabenosyn5 [439-503], EEA1 [36-126], FYCO1 [963–1206], Rim2 [27-175], and Rim1 [20-227] as RBDs, respectively (Simonsen et al., 1998; Pankiv et al., 2010; Fukuda, 2003; Eathiraj et al., 2005). To test the sensitivity and specificity of these Rab sensors, we took three approaches. First, we transfected wild-type (WT)-Rab, dominant-negative (DN)-Rab, and constitutively active (CA)-Rab sensors in HEK 293T cells. As expected, DN-Rab and CA-Rab sensors displayed lower and higher binding fractions than the WT-Rab sensor, respectively, indicating the sensitivity of Rab sensors (Figure 1b and c). Particularly, Rab4 and Rab10 sensors showed significantly different binding fractions between WT- and DN- or CA-Rabs (Figure 1b and c). For the Rab10 sensor, we used the mTurquoise2-mVenus pair instead of the mEGFP-mCherry pair since it reported higher binding fraction differences between WT- and DN-, as well as WT- and CA-Rab10 sensors in HEK 293T cells (Figure 1b,c, Figure 1—figure supplement 1a-d; Takahashi et al., 2015; Goedhart et al., 2012). Second, we coexpressed a Rab sensor with the corresponding Rab GAPs or GEFs to test whether the sensor could respond to the known upstream signaling (Yoshimura et al., 2010; Tall et al., 2001; Rink et al., 2005; Itoh et al., 2006; Hattula et al., 2002; Fukuda et al., 2008; Chamberlain et al., 2004). Indeed, these GAPs and GEFs respectively decreased and increased the activity of Rab proteins as reported by the sensor (Figure 1d–I). Compared with Rab5 and Rab8, Rab4, Rab7, and Rab10 sensors displayed higher binding fraction changes in response to GAPs or GEFs (Figure 1d–i). Third, we measured Rab activity change in response to N-Methyl-D-aspartic acid (NMDA) application in neurons (Murakoshi et al., 2011). We biolistically transfected rat organotypic hippocampal slices with each Rab sensor and imaged the proximal apical dendrites of CA1 pyramidal neurons. Bath application of NMDA (15 µM, 2 min) in zero extracellular Mg²⁺ triggered a robust activation of Rab sensors in the dendrites, suggesting that Rab sensors could report neuronal Rab activities (Figure 1j). Nevertheless, these Rab sensors showed different activation kinetics upon NMDA stimulation (Figure 1j, Figure 1—figure supplement 1e). Rab4 sensor was rapidly activated, peaked at 3 min and subsequently decayed (Figure 1j, Figure 1—figure supplement 1e). On the contrary, Rab5, 7, 8, and 10 sensors displayed a gradually accumulated activation pattern, which peaked at 6–11 min and decreased afterwards (Figure 1j, Figure 1—figure supplement 1e). Notably, the Rab10 sensor was transiently inactivated in the first 2 min, possibly reflecting a fast response to NMDAR activation (Figure 1j, Figure 1—figure supplement 1e). Overall, these results demonstrate that our Rab sensor strategy is generalizable to many Rab proteins.

We biolistically transfected cultured organotypic hippocampal slices of rats with Rab10 sensor and imaged the secondary apical dendrites of CA1 pyramidal neurons. mTurquoise2-Rab10, the sensor donor, was continuously distributed in the dendrites and spines, without showing an endosomal punctate pattern (Figure 2a). To eliminate the effect of protein overexpression on its localization, we probed endogenous Rab10 by CRISPR-Cas9-mediated homology-directed repair (SLENDR) technique in vivo (Figure 1—figure supplement 1f–j; Mikuni et al., 2016). Endogenous Rab10 displayed a heterogeneous pattern: it is widely distributed along the dendrite and protrudes into the spines (Figure 1—figure supplement 1i). To elucidate the subcellular compartmentalization of Rab10, we further examined its colocalization with exogenously expressed endosomal markers in vivo (Figure 1—figure supplement 1j). Rab10 is significantly overlapped with Rab7-labeled lysosomes, partially colocalized with Rab11-labeled recycling endosomes, but separated from Rab5-labeled early endosomes (Figure 1—figure supplement 1j).

To characterize the spatiotemporal dynamics of Rab10 activity in single dendritic spines during sLTP, we combined two-photon glutamate uncaging with 2pFLIM (Murakoshi et al., 2011; Harward et al., 2016; Harvey et al., 2008; Hedrick et al., 2016), and imaged the secondary apical dendrites of CA1 pyramidal neurons expressing the Rab10 sensor at 25–27°C. Upon focal glutamate uncaging (0.5 Hz, 60 s) in zero extracellular Mg²⁺, a single spine underwent a rapid volume increase within the first few minutes (ΔV_transient = 275.9 ± 25.0%; Figure 2a, e and f), which decayed to a smaller but sustained volume increase lasting more than 30 min (ΔV_sustained = 79.6 ± 6.7%; Figure 2a, e and g), consistent with the previous studies (Harvey et al., 2008; Hedrick et al., 2016; Matsuzaki et al., 2004; Murakoshi et al., 2011; Harward et al., 2016). Concomitant with the spine enlargement, Rab10 showed a rapid and spine-specific decrease of activity in the stimulated spines, which lasted for more than 30 min (Figure 2a–d). Neither the changes in spine volume, basal binding fraction, nor the changes in the binding fraction of Rab10 sensor correlated with the initial spine size (Figure 2—figure supplement 1a–e). To verify the specificity of the sensor response, we replaced the RBD of the Rab10 sensor with the RBD of Rab4 sensor (Rabenosyn5 [439-503], false acceptor). This false acceptor sensor did not show a significant change in FRET (measured as the binding fraction change) in the stimulated spines (Figure 2c,d,f,g, Figure 2—figure supplement 2). In addition, the Rab10 sensor displayed a similar inactivation pattern at a near-physiological temperature (33–35°C; Figure 2—figure supplement 3a–d). The inactivation of Rab10 was correlated neither with the concentration of donor nor with that of the acceptor, suggesting that overexpressing Rab10 sensor has minimal effects on Rab10 inactivation (Figure 2—figure supplement 3e–h). In summary, our results demonstrate that Rab10 is persistently inactivated in the stimulated spines during sLTP, and this inactivation is compartmentalized in the stimulated spines.

The persistent inactivation of Rab10 during sLTP may be caused by the dilution of active Rab10 proteins due to the rapid spine enlargement. To examine this possibility, we compared the basal binding fraction of Rab10 sensor in spines and dendrites and analyzed the subset of spines with lower Rab10 basal activity than the dendrite (Figure 2—figure supplement 4b–d). We found that Rab10 activity still decreased against the gradient at the spine neck upon sLTP induction (Figure 2, Figure 2—figure supplement 4d). In addition, the binding fraction of the Rab10 sensor is independent of donor intensity (Figure 2—figure supplement 4a). Thus, instead of passive signal dilution, the persistent inactivation of Rab10 requires active biological processes, presumably by translocating Rab10-positive vesicles out of the spine or directly inactivating Rab10 in the spine. To examine whether Rab10 distribution is changed during sLTP, we further analyzed the intensity of Rab10 donor in the stimulated spines and dendrites. Upon sLTP induction, Rab10 donor intensity significantly increased in the stimulated spines but remained unchanged in the dendrites (Figure 2—figure supplement 5a). The dramatic increase of Rab10 intensity in the stimulated spines was probably due to passive diffusion by spine enlargement. Therefore, we further analyzed the mean intensities of Rab10 donor in the stimulated spines and dendrites, which were similar prior to and post sLTP induction (Figure 2—figure supplement 5b and c).

To further identify signaling pathways that inactivate Rab10 during sLTP, we applied pharmacological inhibitors targeting potential upstream components (Patterson and Yasuda, 2011; Hedrick et al., 2016). Inhibition of NMDARs by 2-amino-5-phosphonopentanoic acid (AP5, 50 µM) completely abolished Rab10 inactivation and spine enlargement (Figure 2c,d,f,g, Figure 2—figure supplement 6a,b). Application of CN21 (10 µM), a CaMKII inhibitory peptide (Chang et al., 1998; Vest et al., 2007), abolished Rab10 inactivation while partially attenuating the volume change (Figure 2c,d,f,g, Figure 2—figure supplement 6c,d). In contrast, mitogen-activated protein kinase (MAPK)/ERK kinase (MEK) inhibitor U0126 (20 µM) had no effect on Rab10 inactivation, although it impaired sLTP during the sustained phase (Figure 2c,d,f,g, Figure 2—figure supplement 6e,f). These results demonstrate that Rab10 is persistently inactivated in the stimulated spines during sLTP, and this inactivation is dependent on NMDARs and CaMKII but not on the MAPK/ERK signaling pathway.

Next, we measured the spatiotemporal profile of Rab4 activity in dendrites during sLTP induced in single spines. Consistent with its localization in early and recycling endosomes (Ehlers, 2000; Sönnichsen et al., 2000; van der Sluijs et al., 1992b), mEGFP-Rab4, the sensor donor, showed a punctate distribution pattern (Figure 3a). In the basal state, Rab4 sensor activity is not correlated with the donor intensity and was lower in the spines than dendrites (Figure 2—figure supplement 4e–g). Glutamate uncaging (0.5 Hz, 60 s) induced a transient and sustained volume increase in the stimulated spines of neurons expressing Rab4 sensor (ΔV_transient = 341.0 ± 29.4% and ΔV_sustained = 77.6 ± 8.0%; Figure 3a and e–g). In contrast to the persistent inactivation of Rab10, Rab4 activity was transiently elevated in the stimulated spines and decayed, with no activity change in the adjacent spines or dendrites (Figure 3a–d). This enhanced spine activity exceeds that of dendrite during sLTP, suggesting that the activation requires active processes (Figure 2—figure supplement 4h). Similarly to Rab10, we examined the intensity and mean intensity of Rab4 donor in the stimulated spines and dendrites. In the transient phase of sLTP, both the intensity and mean intensity of Rab4 donor significantly increased in the stimulated spines, suggesting recruitment of Rab4 into the stimulated spines (Figure 2—figure supplement 5d–f). In contrast, there was no donor intensity or mean intensity change in the dendrites (Figure 2—figure supplement 5d–f). Moreover, the initial spine volume was not correlated either with the volume changes, the basal activity of Rab4 reported or the level of Rab4 activation reported by the sensor (Figure 2—figure supplement 1f–j). Replacing the RBD of Rab4 sensor with the RBD for Rab10 sensor (Rim1 [20-227], false acceptor) showed no change of the binding fraction in the stimulated spines during sLTP, indicating the specificity of the sensor response (Figure 3c,d,f,g, Figure 3—figure supplement 1). In addition, the Rab4 sensor showed a similar transient activation pattern at a near-physiological temperature (33–35°C; Figure 3—figure supplement 2). The obtained signal was independent of the expression level of Rab4 sensor (Figure 3—figure supplement 2e–h). Thus, our results demonstrate that Rab4 is transiently activated in the stimulated spines during sLTP, and this activation is compartmentalized in the stimulated spines.

To identify signaling pathways that activate Rab4 during sLTP, we applied pharmacological inhibitors targeting putative upstream components. Inhibition of NMDARs by AP5 (50 µM) completely abolished Rab4 activation and spine enlargement (Figure 3c,d,f,g, Figure 3—figure supplement 3a,b). Application of CN21 (10 µM) decreased Rab4 activity and volume changes both in the transient phase and sustained phase (Figure 3c,d,f,g, Figure 3—figure supplement 3c,d). In contrast, MEK inhibitor U0126 (20 µM) had no effect on Rab4 activation, although it impaired sLTP during the sustained phase (Figure 3c,d,f,g, Figure 3—figure supplement 3e,f). Altogether, Rab4 activation during sLTP is dependent on NMDARs and CaMKII, but not on the MAPK/ERK signaling pathway.

Given that Rab10 and Rab4 display opposing activity profiles during sLTP, we hypothesized that they would have opposite functions in spine structural plasticity. To test this hypothesis, we knocked down Rab10 or Rab4 by respective shRNA and examined the effects of these manipulations on sLTP. We biolistically transfected cultured organotypic hippocampal slices of rats with scrambled shRNA control or shRNA against Rab10 or Rab4, together with mEGFP as the volume marker. Compared with scrambled shRNA control, knockdown of Rab10 had no effect on spine size but increased spine density (Figure 4a, Figure 4—figure supplement 1a,b,f,g,h,i). Knockdown of Rab4 had no effect on spine size or density (Figure 4a, Figure 4—figure supplement 1a,b,e,g,h,i).

We further induced sLTP by two-photon glutamate uncaging in single spines of neurons expressing mEGFP (Murakoshi et al., 2011; Lee et al., 2009). Under control conditions with scrambled shRNA, application of a train of glutamate uncaging pulses (0.5 Hz, 60 s) in zero extracellular Mg²⁺ induced a rapid spine volume increase in the transient phase, which decayed to a sustained enlarged volume for more than 30 min (Figure 4a and c). However, knockdown of Rab10 by shRNA enhanced spine enlargement both in the transient and sustained phase of sLTP, which was rescued by co-expressing shRNA-resistant Rab10 (for scrambled shRNA, ΔV_transient = 215.5 ± 16.6% and ΔV_sustained = 64.4 ± 9.9%; for Rab10 shRNA, ΔV_transient = 309.0 ± 36.3% and ΔV_sustained = 140.6 ± 16.4%; for Rab10 rescue, ΔV_transient = 213.5 ± 19.7% and ΔV_sustained = 87.4 ± 11.7%; Figure 4a,c,e, Figure 4—figure supplement 1a,b,d,f,g). In contrast, knockdown of Rab4 by shRNA significantly impaired the transient phase of sLTP while leaving the sustained phase intact (for scrambled shRNA, ΔV_transient = 291.6 ± 35.6% and ΔV_sustained = 76.1 ± 10.0%; for Rab4a/4b shRNA, ΔV_transient = 119.8 ± 15.4% and ΔV_sustained = 73.8 ± 12.1%; Figure 4a,c,e, Figure 4—figure supplement 1a,b,e,g). This phenotype was rescued by coexpression of shRNA-resistant Rab4a (ΔV_transient = 223.1 ± 32.8% and ΔV_sustained = 84.2 ± 12.3%; Figure 4a,c,e, Figure 4—figure supplement 1c). Overall, these results suggest that Rab10 negatively regulates both the transient and sustained phases of sLTP, while Rab4 is required for the transient phase of sLTP.

As an alternative strategy to inhibit Rab10 and Rab4 functions, we examined the effects of overexpressing DN-Rab mutants on spine structural plasticity. Consistent with the shRNA results, DN-Rab10 enhanced both the transient and sustained phase of sLTP (for control, ΔV_transient = 209.2 ± 31.1% and ΔV_sustained = 64.8 ± 9.4%; for Rab10 DN, ΔV_transient = 332.0 ± 31.8% and ΔV_sustained = 125.0 ± 17.4%; Figure 4b, d and f), while DN-Rab4a selectively inhibited the transient phase of sLTP (for control, ΔV_transient = 357.4 ± 52.0% and ΔV_sustained = 83.5 ± 10.8%; for Rab4a DN, ΔV_transient = 198.1 ± 32.1% and ΔV_sustained = 70.8 ± 11.0%; Figure 4b, d and f). Moreover, we evaluated the effect of overexpressing CA-Rab10 or CA-Rab4 on sLTP. These manipulations in general caused opposite phenotypes to the DN mutants: CA-Rab10 decreased both the transient and sustained phase of sLTP (for control, ΔV_transient = 209.2 ± 31.1% and ΔV_sustained = 64.8 ± 9.4%; for Rab10 CA, ΔV_transient = 97.7 ± 16.5% and ΔV_sustained = 23.0 ± 7.1%; Figure 4b, d and f), while CA-Rab4 slightly increased the transient phase of sLTP (but not statistically significant; for control, ΔV_transient = 357.4 ± 52.0% and ΔV_sustained = 83.5 ± 10.8%; for Rab4a CA, ΔV_transient = 463.1 ± 61.1% and ΔV_sustained = 112.7 ± 18.3%; Figure 4b, d and f).

We further evaluated the effects of Rab10 deletion on structural and electrophysiological LTP using Rab10 conditional knockout mice (Rab10^fl/fl) (Vazirani et al., 2016). For sLTP measurement, we biolistically transfected cultured organotypic hippocampal slices of Rab10^fl/fl mice with tdTomato-fused Cre recombinase and mEGFP, or tdTomato and mEGFP as a control (J.-Y. Chang et al., 2017). Consistent with the Rab10-knockdown results, deletion of the Rab10 gene increased sLTP in the stimulated spines of CA1 pyramidal neurons (Figure 4g, h and i). Furthermore, we crossed Rab10^fl/fl mice with Camk2a-Cre mice (Rab10^fl/fl:Camk2a-Cre^+/-) to postnatally remove Rab10 from forebrain excitatory neurons (Tsien et al., 1996). These animals showed enhanced LTP upon theta burst stimulation (TBS) at Schaeffer collateral synapses (Figure 4k and l), with the basal synaptic transmission unchanged (Figure 4j). Control animals (Rab10^fl/fl without Cre) showed a modest LTP under this condition (Figure 4k and i; Gong et al., 2009; Grover et al., 2009; Capocchi et al., 1992). Moreover, we monitored Schaffer collateral synaptic transmission in these mice and found no difference in the amplitude of AMPAR- and NMDAR-EPSCs, or AMPAR/NMDAR EPSC ratio (Figure 4—figure supplement 2). These results indicate that Rab10 is a negative regulator for electrophysiological LTP.

Overall, our results demonstrate that Rab10 negatively regulates both the transient and sustained phase of sLTP, while Rab4 positively regulates the transient phase of sLTP. These functions are consistent with the direction and time window of their activity changes during sLTP. Moreover, Rab10 negatively modulates electrophysiological LTP.

Plasmids developed in this study are available at Addgene. Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact Ryohei Yasuda (ryohei.yasuda@mpfi.org).

HEK 293T cells (Fisher Scientific) were grown in DMEM (Gibco) supplemented with 10% fetal bovine serum (Invitrogen) and 1% penicillin-streptomycin (Invitrogen). The cell cultures were maintained at 37°C in a 5% CO₂ humidified atmosphere.

CD wild-type rats (CD IGS) purchased from Charles River Laboratories were used for preparation of hippocampal organotypic slice culture and dissociated postnatal cortical neuron culture. Both male and female animals were used and randomly allocated to experimental groups. All the experiments were performed in accordance with guidelines from the US National Institutes of Health and were approved by Duke University Medical Center and the Institutional Animal Care and Use Committee of Max Planck Florida Institute for Neuroscience.

Rab10 conditional knockout (Rab10^fl/fl) mouse was a gift from Dr. Timothy E McGraw (Vazirani et al., 2016). Camk2a-Cre mice were previously reported (Tsien et al., 1996). Swiss Webster mice were obtained from Charles River for endogenous Rab10 knockin experiments. Both male and female mice were used. All the experiments were performed in accordance with guidelines from the US National Institutes of Health and were approved by the Institutional Animal Care and User Committee of Max Planck Florida Institute for Neuroscience.

Organotypic rat or mouse hippocampal slices were prepared at postnatal day 6 or 7, as previously described (Stoppini et al., 1991). Briefly, coronal hippocampal slices were dissected at 400 μm thickness using a McIlwain tissue chopper (Ted Pella, Inc). The slices were cultured on hydrophilic PTFE membranes (Millicell, Millipore), which were inserted in the culture medium containing 8.4 mg/ml MEM (Sigma), 20% horse serum (Gibco), 1 mM L-Glutamine (Sigma), 5.2 mM NaHCO₃, 12.9 mM D-Glucose, 0.075% Ascorbic acid, 30 mM Hepes,1 µg/ml Insulin, 1 mM CaCl₂ and 2 mM MgSO₄. The slice cultures were maintained at 35 °C in a 5% CO₂ humidified atmosphere.

Dissociated postnatal cortical cultures were prepared as previously published (Shibata et al., 2015). Briefly, cortices dissected from newborn rats (random male and female) were triturated and plated into 5 cm dishes coated with 50 µg/ml PLL (Sigma) in culture medium consisting of Basal Medium Eagle (BME) supplemented with 10% heat-inactivated fetal bovine serum (Invitrogen), 35 mM glucose (Sigma), 1 mM L-glutamine (Sigma), 100 U/ml penicillin (Sigma), and 0.1 mg/ml streptomycin (Sigma). Neuron cultures were maintained at 35°C in a 5% CO₂ humidified atmosphere.

To generate pCAGGS-mCherry, pCAGGS-mTurquoise2, pCAGGS-mCherry-mCherry, and pCAGGS-mVenus-mVenus, the respective fluorescence protein sequence was cloned into pCAGGS backbone from Raichu-2517KX gifted from Dr. Michiyuki Matsuda (Kitano et al., 2008). Rat full-length Rab4a, Rab5a, Rab7a, Rab8a, Rab11a, and Rab10 were amplified by PCR from rat brain cDNA library (Dharmacon, Cat# LRN1205) and cloned into pmEGFP-C1 (Murakoshi et al., 2011), pCAGGS-mEGFP, pCAGGS-mCherry, and pCAGGS-mTurquoise2. All FRET donors were tagged at the amino terminus of Rab proteins. The linker between FRET donors (mEGFP or m Turquoise2) and Rab GTPases is SGLRSRG. Rabenosyn5 [439-503], EEA1[36-126], FYCO1[963–1206], Rim2 [27-175], and Rim1 [20-227] cDNAs were amplified by PCR from rat brain cDNA library and inserted into pCAGGS-mCherry-mCherry and pCAGGS-mVenus-mVenus constructs. The second mVenus has a mutation L68V, which improves bleaching time by 30% (called SYFP2; Kremers et al., 2006). The linkers between mCherry and RBDs are SGLRSRA for the amino terminus and GSG for the carboxy terminus. The linkers between mVenus and Rim1 [20-227] are SGLRSRG for the amino terminus and GSG for the carboxy terminus. Rab dominant negative (DN) and constitutively active (CA) mutants were generated from wild type Rab GTPases by site-directed mutagenesis and subcloned into pCAGGS-mEGFP, pCAGGS-3Flag, and pCAGGS-mCherry constructs. PIK3R1 [114-313], Rin1, RNTre (USP6NL), RabGAP5 (SGSM3), GRAB (Rab3il1), Rabin8 (Rab3ip), TBC1D30, Evi5l, and VPS39 were amplified from rat brain cDNA library and cloned into pCAGGS or pCAGGS-3HA vector. Full-length Dennd4c, TBC1D15, and Rab3gap1 were amplified by PCR from MGC mouse cDNA (Dharmacon) and subcloned into pCAGGS-3HA construct. psiCHECK-2-Sal4-wt_3'UTR was a gift from Robert Blelloch (Addgene plasmid # 31862). psiCHECK-2-Rab GTPases were generated by inserting Rab GTPases into psiCHECK-2-Sal4-wt_3'UTR by XhoI/NotI.Tetanus toxin light chain (Eisel et al., 1993) was subcloned into pCAGGS-3Flag construct. SEP-GluA1 was a gift from Dr. Scott Soderling at Duke University (Wang et al., 2008). mTurquoise2-pBAD and mVenus-pBAD were gifts from Michael Davidson (Addgene plasmid # 54844 and # 54845). The human codon-optimized S. pyogenes Cas9 (SpCas9) and single guide RNA (sgRNA) expression plasmid was a gift from F. Zhang (pX330, Addgene plasmid # 42230; Cong et al., 2013).

These antibodies were used for SDS-PAGE and immunoblotting: rabbit anti-Rab10 (1:500; Cell Signaling Technology, #8127), rabbit anti-Rab4b (1:500; Thermo Fisher Scientific, #PA5-49124), mouse anti-Rab4a (1:500; ThermoFisher Scientific, #MA5-17161) and mouse anti-β-actin (1:2000; Sigma, #A5316), HRP-conjugated goat anti-rabbit (1:5000, Bio-Rad, #170–6515) and HRP-conjugated goat anti-mouse (1:5000, Bio-Rad, #172–1011). These antibodies were used for histology: rabbit anti-HA primary antibody (1:1000, Cell Signaling Technology, #3724), Goat anti-Rabbit IgG (H+L) Secondary Antibody, Alexa Fluor 568 conjugate (1:1000, Thermo Fisher Scientific, #A-11036).

For shRNA-mediated knock-down of Rab4 and Rab10, we used SHCLND-NM_009003 plasmid for Rab4a (Sigma-Aldrich, TRCN0000088975), SHCLND-NM_016154 plasmid for Rab4b (Sigma-Aldrich, TRCN0000380038), and TRC-Mm1.0 plasmid for Rab10 (Dharmacon, TRCN0000100838). The respective shRNA sequences (according to manufacturer and sequencing confirmation) are CCGGAGATGACTCAAATCATACCATCTCGAGATGGTATGATTTGAGTCATCTTTTTTG for Rab4a, GTACCGGGGTCATCCTCTGTGGCAACAACTCGAGTTGTTGCCACAGAGGATGACCTTTTTTG for Rab4b, and TTGCCTTTCGGTACAACTCTC (mature antisense) for Rab10. For shRNA control, we used scrambled shRNA with the following sequence: CCTAAGGTTAAGTCGCCCTCGCTCGAGCGAGGGCGACTTAACCTTAGG (Addgene plasmid # 1864). To visualize transfected neurons in sLTP experiments, mEGFP was inserted into scrambled shRNA, Rab4a and Rab10 shRNA by KpnI/BamHI, and Rab4b shRNA by BamHI/BstEII. The mEGFP expression was driven by a separate hPGK promoter (shRNA/mEGFP). For the rescue experiments, silent mutations of three amino acids were introduced at the targeted region for Rab4a and Rab10 by site-directed mutagenesis (for shRNA-resistant Rab4a, AAAGATGACTCCAACCACACCATA; for shRNA-resistant Rab10, GAGAGTTGTGCCCAAGGGCAA).

HEK 293T cells were transfected with Lipofectamine 2000 following the manufacturer’s recommendations (Invitrogen), and imaged 24–48 hr after transfection. For Rab GTPase FRET sensors, the ratio of transfected FRET donor and acceptor was 1:3.

After 9–13 days in culture, organotypic hippocampal slices were transfected biolistically with gene gun (McAllister, 2000; Bio-Rad, pressure 200 psi) using gold beads (Bio-Rad, 1.6 µm) coated with plasmids, and imaged 3–4 days after transfection. For the Rab4 FRET sensor, pmEGFP-Rab4a and pCAGGS-mCherry-Rabenosyn5 [439-503]-mCherry (1:1, 20 µg) were expressed for 3 days. For the Rab10 FRET sensor, pCAGGS-mTurquoise2-Rab10 and pCAGGS-mVenus-Rim1 [20-227]-mVenus (1:3, 40–60 µg) were expressed for 3–4 days.

We used a custom-built two-photon fluorescence lifetime imaging microscope (2pFLIM) with two Ti:Sapphire lasers (Chameleon, Coherent) as previously described (Murakoshi et al., 2008; Yasuda, 2006). One laser was tuned to 920 nm to excite both donor for lifetime measurement and acceptor for morphology. The second laser was tuned to 720 nm for glutamate uncaging. The imaging power for two lasers was controlled independently by electro-optical modulators (Conoptics). The fluorescence was collected by an objective (60 X, 1.0 numerical aperture, Olympus), separated by a dichroic mirror (Chroma, 565 nm for mEGFP/mCherry and 505 nm for mTurquoise2/mVenus), filtered by wavelength filters (Chroma, ET520/60 M-2p for mEGFP, ET620/60 M-2p for mCherry, ET480/40 M-2p for mTurquoise2, ET535/50 M-2p for mVenus), and finally detected by two independent photoelectron multiplier tubes (PMTs). We used 1.2–1.5 mW imaging power for mEGFP/mCherry sensor, and 1.6–1.8 mW for mTurquoise2/mVenus sensor.

Two-photon fluorescence lifetime imaging in HEK 293T cells was performed in imaging solution containing 20 mM HEPES (pH 7.3), 130 mM NaCl, 2.5 mM KCl, 2 mM MgCl₂, 2 mM NaHCO₃, 1.25 mM NaH₂PO₄ and 25 mM D-glucose.

Two-photon lifetime imaging and glutamate uncaging in organotypic slices was performed in Mg²⁺-free artificial cerebrospinal fluid (ACSF; 127 mM NaCl, 2.5 mM KCl, 1.25 mM NaH₂PO₄, 25 mM NaHCO₃, 25 mM D-glucose, aerated with 95% O₂ and 5% CO₂) with 4 mM CaCl₂, 4 mM MNI-caged glutamate (Tocris) and 1 μM tetrodotoxin (TTX, Enzo). Uncaging pulses (0.5 Hz, 60 s, 4–6ms, 3.5–3.8 mW) were delivered to the back focal aperture of the objective, which was around 0.5 µm from the tip of the spine head. The adjacent spine refers to the first or second spine located next to the stimulated spine, typically positioned opposite to the stimulated spine. Additionally, the size of the adjacent spine must be sufficiently large for imaging. We used a heater controller (Warner Instruments TC-344B) to monitor the temperature at 25–27°C or 33–35°C. Images were analyzed by MATLAB (MathWorks) and ImageJ.

The control and experimental groups were derived from the same batch of samples and were imaged on the same day to ensure consistent microscope settings and imaging buffer conditions, but in an unblinded manner. For the AP5 experiment, control samples were incubated in the imaging buffer for 30 min prior to the induction of sLTP. Following the control experiments, AP5 was added to the same imaging buffer, and samples were incubated for an additional 30 min before sLTP induction. In the CN21 and U0126 experiments, scrambled peptide or U0124 (as controls) was added into the imaging buffer, and samples were incubated for 30 min prior to sLTP induction. After the control experiments, the perfusion system was thoroughly washed with imaging buffer. For the experimental groups, CN21 peptide or U0126 was then added to the imaging buffer, and samples were incubated for 30 min before sLTP induction.

As described previously (Harvey et al., 2008), the fraction of donor bound to acceptor was determined by fitting a fluorescence lifetime curve that summed contributions from all pixels in the image, using a double exponential function convolved with the Gaussian pulse response function, as shown in the equation:F(t)=F0[PDH(t,t0,τD,τG)+PADH(t,t0,τAD,τG)]\begin{document}$$\displaystyle F(t) = F_0 \left[ P_DH(t,t_0,\tau_D,\tau_G)+P_{AD}H(t,t_0,\tau_{AD}, \tau_G)\right]$$\end{document}

In this equation, F₀ is the peak fluorescence before convolution, while PD\begin{document}$P_{D}$\end{document} and PAD\begin{document}$P_{AD}$\end{document} represent the fractions of free donor and donor bound to the acceptor, respectively. t₀ is the time offset, τD\begin{document}$\tau _{D}$\end{document} is the fluorescence lifetime of the free donor, τAD\begin{document}$\tau _{AD}$\end{document} is the fluorescence lifetime of donor bound with acceptor, τG\begin{document}$\tau_{G}$\end{document} defines the width of the Gaussian pulse response function, and H(t) is a fluorescence lifetime curve with a single exponential function convolved with the Gaussian pulse response function:H(t,t0,tD,tG)=12exp⁡(τG22τD2−t−t0τD)erfc(τG2−τD(t−t0)2τDτG)\begin{document}$$\displaystyle H\left (t,t_0,t_D,t_G\right)=\frac{1}{2}\exp\left(\frac{\tau_G^2}{2\tau_D^2}-\frac{t-t_0}{\tau _D}\right)erfc\left(\frac{\tau_G^2-\tau_D\left(t-t_0\right)}{\sqrt2\tau_D\tau_G}\right) $$\end{document}

in which erfc is the complementary error function. We fixed τD\begin{document}$\tau_{D}$\end{document} as 2.46 ns, 2.60 ns, and 4.15 ns, corresponding to the free mEGFP–Rab4a, mEGFP-Rab10, and mTurquoise2-Rab10 donors, respectively. Similarly, τAD\begin{document}$\tau_{AD}$\end{document} was set to the fluorescence lifetime of the donor when bound to its acceptor, with values of 1.10 ns for the mEGFP/mCherry pair and 1.60 ns for the mTurquoise2/mVenus pair.

To create the fluorescence lifetime image, we computed the mean photon arrival time, <t>, for each pixel using the following equation:<t>=∫tF(t)dt/∫F(t)dt\begin{document}$$\displaystyle \lt t\gt = \int tF\left (t\right)dt/\int F\left (t\right)dt$$\end{document}

The mean photon arrival time is then related to the mean fluorescence lifetime, <τ>, by an offset arrival time, t0\begin{document}$t_{0}$\end{document}, which is determined through image-wide fitting:<τ>=<t>− t0\begin{document}$$\displaystyle \lt \tau \gt = \lt t\gt -\ t_{0}$$\end{document}

For small regions of interest (ROIs) within an image, such as spines or dendrites, the binding fraction (P_AD) was calculated using the formula:PAD=τD(τD−<τ>)(τD−τAD)(τD+τAD−<τ>)\begin{document}$$\displaystyle P_{AD}=\frac{\tau _{D}\left (\tau _{D}-\lt \tau \gt \right)}{\left (\tau _{D}-\tau _{AD}\right)\left (\tau _{D}+\tau _{AD}-\lt \tau \gt \right)}$$\end{document}

To estimate the spine volume in neurons expressing Rab sensors, we measured the integrated fluorescence intensity of mCherry-RBD-mCherry or mVenus-RBD-mCherry in the spine, which is proportional to the spine volume (Holtmaat et al., 2005), and normalized it by the fluorescence intensity in the thick apical dendrite from the same neuron. We further multiplied this normalized value by the volume of the point spread function, which gives the spine volume in fL (Nimchinsky et al., 2004; Harvey et al., 2008).

For the sensor experiments, we used mCherry as a volume indicator. There is significant bleed-through of donor fluorescence into the acceptor channel (Yasuda, 2006). However, we anticipate that this would not have a large impact on the estimation of spine volume changes, as fluorescence changes in both the red and green channels exhibit similar time courses (Figures 2e and 3e, Figure 2—figure supplements 5a and 6d).

Rab4 is located outside of the endosome (van der Sluijs et al., 1991), making it unlikely that intra-vesicular pH changes affect fluorescence lifetime measurements. The fluorescence lifetime of EGFP is minimally pH-dependent, as its protonated state does not fluoresce (Kneen et al., 1998). Additionally, mCherry is highly resistant to pH variations, with a pKa of less than 4.5 (Shaner et al., 2004). Therefore, we conclude that endosomal pH changes are unlikely to impact our Rab activity measurements.

Rat organotypic hippocampal slices (DIV 9-DIV 13) were biolistically transfected with indicated Rab sensors. After 3–4 day expression, CA1 pyramidal neurons were imaged in the basal solution (ACSF with 2 mM CaCl₂, 2 mM MgCl₂ and 1 μM TTX) for 6 min. NMDA (Tocris) was bath-applied in the zero Mg²⁺ solution (ACSF with 4 mM CaCl₂, 15 µM NMDA and 1 μM TTX) for 2 min, and replaced by the washout solution (ACSF with 2 mM CaCl₂, 2 mM MgCl₂, 1 μM TTX and 50 µM AP5) for 32 min.

Rat organotypic hippocampal slices were biolistically transfected with indicated constructs at days in vitro 9–13 (DIV 9-DIV 13). The constructs for Rab shRNA knockdown experiments were: scrambled shRNA/mEGFP (Ctrl shRNA); Rab4a shRNA/mEGFP and Rab4b shRNA/mEGFP (Rab4a/4b shRNA); Rab4a shRNA/mEGFP, Rab4b shRNA/mEGFP, and pCAGGS-mCherry-shRNA-resistant Rab4a (Rab4a rescue); Rab10 shRNA/mEGFP (Rab10 shRNA); Rab10 shRNA/mEGFP and pCAGGS-mCherry-shRNA-resistant Rab10 (Rab10 rescue). The constructs for DN- and CA-Rab overexpression experiments were: pCAGGS-mEGFP (Ctrl); pCAGGS-mEGFP and pCAGGS-mCherry-Rab DN (Rab DN); pCAGGS-mEGFP and pCAGGS-mCherry-Rab CA (Rab CA). The constructs were expressed for 4–5 days for Rab shRNA knock-down and 2–3 days for DN- or CA-Rab mutant overexpression. CA1 pyramidal neurons were imaged in ACSF (aerated with 95% O₂ and 5% CO₂) with 4 mM CaCl₂, 4 mM MNI-caged glutamate and 1 μM TTX. Two-photon glutamate uncaging (0.5 Hz, 60 s, 4–6ms, 3.5–3.8 mW) was performed at a single spine. All experiments were paired with the same day controls from the same batch of slices. The acquired images were analyzed by MATLAB (MathWorks).

Organotypic hippocampal slices of Rab10^fl/fl mice were biolistically transfected with tdTomato-Cre and mEGFP, or tdTomato and mEGFP as a control at DIV 10. At DIV 14, CA1 pyramidal neurons were imaged in ACSF (aerated with 95% O₂ and 5% CO₂) with 4 mM CaCl₂, 4 mM MNI-caged glutamate, and 1 μM TTX. Two-photon glutamate uncaging (0.5 Hz, 60 s) was performed at a single spine. All experiments were paired with the same day controls from the same batch of slices. The acquired images were analyzed by MATLAB (MathWorks).

Rat organotypic hippocampal slices were biolistically transfected with indicated constructs at DIV 9. The constructs for Rab shRNA knockdown experiments were: scrambled shRNA/mEGFP (Ctrl shRNA), Rab4a shRNA/mEGFP and Rab4b shRNA/mEGFP (Rab4a/4b shRNA), or Rab10 shRNA/mEGFP (Rab10 shRNA). After 4 days of expression, CA1 pyramidal neurons were imaged in ACSF by two-photon microscopy at 25–27°C. All experiments were paired with the same-day controls from the same batch of slices. The acquired images were analyzed by MATLAB (MathWorks) and ImageJ.

Rat organotypic hippocampal slices were biolistically transfected with indicated constructs at DIV 9-DIV 13. The DNA constructs for each condition were: pCAGGS-mCherry, SEP-GluA1, and scrambled shRNA (Ctrl shRNA); pCAGGS-mCherry, SEP-GluA1, and pCAGGS-3Flag-TeTxLC (TeTxLC); pCAGGS-mCherry, SEP-GluA1, Rab4a shRNA, and Rab4b shRNA (Rab4a/4b shRNA); pCAGGS-mCherry, SEP-GluA1, and Rab10 shRNA (Rab10 shRNA); pCAGGS-mCherry, SEP-GluA1, Rab4a shRNA, Rab4b shRNA, and pCAGGS-3Flag-shRNA-resistant Rab4a (Rab4a rescue); pCAGGS-mCherry, SEP-GluA1, Rab10 shRNA, and pCAGGS-3Flag-shRNA-resistant Rab10 (Rab10 rescue). After 4 days expression, CA1 pyramidal neurons were imaged in ACSF (aerated with 95% O₂ and 5% CO₂) with 4 mM CaCl₂, 4 mM MNI-caged glutamate, and 1 μM TTX at 25–27°C. All experiments were paired with the same day controls from the same batch of slices. After taking five baseline images of a secondary apical dendrite (1 min interval) with 1.2–1.5 mW imaging laser power, we bleached the whole dendrite by increasing the imaging laser power to 4.0–4.5 mW for 2 min. We then performed two-photon glutamate uncaging (0.5 Hz, 60 s, 4–6ms, 3.5–3.8 mW) at a single spine and collected eight images (1 min interval). The acquired images were analyzed by MATLAB (MathWorks) and ImageJ.

In Figure 5—figure supplement 2, we addressed the impact of SEP-GluA1’s lateral diffusion by subtracting the change in spine surface area (Volume^2/3) rather than correcting for spine volume. This approach is necessary because SEP-GluA1 is only fluorescent on the cell surface, while the fluorescence intensity of cytosolic proteins such as mCherry is proportional to spine volume. Therefore, the overall fluorescence change (ΔF) should be the addition of the contribution from AMPAR trafficking (ΔF_t) and the change in surface area (ΔS) multiplied by the remaining SEP-GluA1 fluorescence per unit area (f):ΔF=ΔFt+fΔS\begin{document}$$\displaystyle \Delta F = \Delta F_t + f \Delta S$$\end{document}

Since fluorescence immediately after photobleaching (before AMPAR trafficking happens), F_o, is given by fS (S is the surface area of the spine):ΔFF0=ΔFtF0+fΔSfS\begin{document}$$\displaystyle \frac{\Delta F}{F_0} = \frac{\Delta F_t}{F_0} + \frac{f \Delta S}{f S} $$\end{document}=ΔFtfS+ΔSS\begin{document}$$\displaystyle = \frac{\Delta F_t}{f S} + \frac{\Delta S}{S} $$\end{document}

Assuming that the surface area change (ΔS/S) is the volume change (ΔV/V) to the power of 2/3, the contribution of the AMPAR trafficking can be calculated as:ΔFtF=ΔFF−(ΔVV)2/3\begin{document}$$\displaystyle \frac{\Delta F_t}{F} = \frac{\Delta F}{F} - \left( \frac{\Delta V}{V} \right)^{2/3} $$\end{document}

Thus, we obtained the contribution of AMPAR trafficking by subtracting the volume change (ΔV/V) to the power of 2/3 from the fluorescence recovery (ΔF/F).

The SLENDR technique was as previously described (Mikuni et al., 2016). Briefly, Rab10 sgRNA (IDT) was ligated into the sgRNA scaffold of pX330 (Ran et al., 2013) to generate Rab10 sgRNA-expressing plasmid. Rab10 sgRNA-expressing plasmid and single-stranded oligodeoxynucleotides (ssODNs; IDT) for homology-directed repair (HDR) were transfected into Neuro 2 a cells by electroporation. Genomic DNA from the electroporation-transduced Neuro 2 a cells was isolated with DNeasy Blood & Tissue Kit (QIAGEN) following the manufacturer’s instruction. Genomic PCR was performed using extracted DNA as a template with corresponding primer set. The PCR product was purified by QiaQuick gel extraction kit (QIAGEN) and then proceeded to Sanger sequencing to detect 2XHA insertion.

In utero electroporation (IUE) was performed as previously described. Electroporation was performed at E13-E14 in anesthetized mice. For endogenous Rab10 localization, 1–2 μl mixture of plasmids (pX330-Rab10 sgRNA and pPB-CAG-mEGFP, 1 μg/μl) and ssODNs (20 μM) was injected into the lateral ventricle of each pup. For colocalization examination of endogenous Rab10 and endosomal markers, 1–2 μl mixture of plasmids (pX330-Rab10 sgRNA and pCAGGS-mEGFP-Rab5a, pCAGGS-mEGFP-Rab7a or pCAGGS-mCherry-Rab11a, 1 μg/μl) and ssODNs (20 μM) was injected into the lateral ventricle of each pup.

With ketamine-xylazine anesthesia (100 μg of ketamine –10 μg of xylazine per g of body weight, i.p.), mice were perfused with 4% paraformaldehyde in 0.1 M phosphate buffer, pH 7.4, and the brain was fixed for 4–12 hr. After washing with PBS, coronal vibratome sections (50 μm in thickness) were prepared (VT1200, Leica). For immunohistochemistry, slices were permeabilized with 0.3–0.4% Triton X-100 in PBS, blocked with 5% normal goat serum and 2% BSA or 5% normal donkey serum in PBS, and incubated overnight with rabbit anti-HA primary antibody (1:1000, Cell Signaling Technology). After 1–3 hr incubation with Alexa Fluor-conjugated secondary antibodies (Invitrogen) followed by DAPI staining (0.1 μg/ml, Life Technologies), slices were imaged using a confocal laser-scanning microscope (LSM880 with Airyscan, Zeiss). Images were processed and analyzed using the Zen (Zeiss) and the Fiji software.

HEK 293T cells were plated into 24-well plates and cotransfected with psiCHECK-2-Rab GTPases and the respective shRNA at a 1:3 ratio. For positive control, we used shRNA against hRluc with the following sequence: TCATAGTAGTTGATGAAGGAG (mature antisense). After 48 hr transfection, luciferase activity was measured using the Dual-Luciferase Reporter Assay System (Promega) following the manufacturer’s protocol. After removing the culture medium, cells were briefly rinsed in pre-warmed 0.1 M phosphate-buffered saline (PBS) and lysed in 100 µL of 1 X passive lysis buffer in the luciferase assay kit. After gently shaking for 15 min at room temperature, samples were prepared in a 96-well plate for luminescence measurement using the GloMax-Multi Detection System (Promega). For data analysis, the hRluc (firefly luciferase) luminescence was normalized by the hluc+ (Renilla Luciferase) luminescence in each well to control for transfection efficiency. All experiments were paired with the same day controls from the same batch of HEK 293T cells.

Dissociated postnatal cortical cultures were prepared as previously published (Shibata et al., 2015). Both male and female animals were used and randomly allocated to experimental groups. Proliferation of non-neuronal cells was inhibited by adding Cytosine arabinoside (2.5 μM) at DIV 2. At DIV 6, cultures were infected with lentiviral particles containing Rab4a shRNA/mEGFP (Rab4a shRNA); Rab4b shRNA/mEGFP (Rab4b shRNA); Rab4a shRNA/mEGFP and Rab4b shRNA/mEGFP (Rab4a/4b shRNA); Rab10 shRNA/mEGFP (Rab10 shRNA) or scrambled shRNA/mEGFP (Ctrl shRNA). At DIV 17, cells were washed with ice-cold PBS and immediately extracted with ice-cold T-PER protein extraction buffer (Pierce) supplemented with inhibitors for proteases and phosphatases (Roche). The lysates were centrifuged at 15,000 × g for 15 min at 4 °C and the supernatants were used for further analysis.

Samples were prepared for standard SDS-PAGE and separated on 12% acrylamide gel (Mini-PROTEAN TGX precast gels, Bio-Rad), then transferred onto 0.2 µm pore size PVDF membranes (Millipore) using semi-dry immunoblotting (transfer buffer containing 25 mM Tris, 200 mM glycine and 20% methanol). Membranes were blocked with 5% nonfat milk (Great Value) in TBS-T (Tris Buffered Saline with 0.1% Tween-20) for 1 hr at room temperature, then incubated overnight at 4°C with primary antibodies diluted in 5% BSA in TBS-T. We used the following commercially available antibodies: rabbit anti-Rab10 (1:500; Cell Signaling Technology), rabbit anti-Rab4b (1:500; Thermo Fisher Scientific), mouse anti-Rab4a (1:500; Thermo Fisher Scientific), and mouse anti-β-actin (1:2000; Sigma). Membranes were washed three times for 15 min in TBS-T, followed by incubation for 2 hr at room temperature with HRP-conjugated goat anti-rabbit or goat anti-mouse secondary antibodies (Bio-Rad), diluted 1:5000 in 5% nonfat milk in TBS-T. Membranes were washed 3 times for 15 minutes in TBS-T, then incubated with Pierce ECL Plus western blotting substrate (for Rab10, Rab4a, and Rab4b) or Pierce ECL western blotting substrate (for β-actin) for detection of western blotted proteins. We used the Image Quant LAS4000 Imaging System (GE Healthcare) to visualize protein bands.

Rab 10 Cre – and Cre +littermate mice (P 30 P 50, blind-coded) were anesthetized by isoflurane inhalation and perfused intracardially with a chilled choline chloride solution. The brain was removed and placed in the same choline chloride solution composed of 124 mM Choline Chloride, 2.5 mM KCl, 26 mM NaHCO₃, 3.3 mM MgCl₂, 1.2 mM NaH₂PO₄, 10 mM Glucose, and 0.5 mM CaCl₂, pH 7.4 equilibrated with 95% O₂ and 5% CO₂. Coronal slices (300 µm) containing the hippocampus were cut using a vibratome (Leica) and maintained in a submerged chamber at 32 °C for 1 hr and then at room temperature in oxygenated ACSF.

All animals were coded for their genotypes, and all recordings and analyses were conducted blindly. Slices were perfused with oxygenated ACSF containing 2 mM CaCl₂, 2 mM MgCl₂, and 100 µM picrotoxin. One glass electrode (resistance ~4 MΩ) containing the same ACSF solution was placed in the dendritic layer of the CA1 area (~100–200 µm away from the soma) while stimulating Schaffer Collateral fibers with current square pulses (0.1ms) using a concentric bipolar stimulation electrode (FHC). The initial slope of the fEPSP was monitored with custom software (Matlab). The stimulation strength was set to ~50% saturation. A 20 min stable baseline was first recorded before induction of LTP. LTP was induced by applying 5 trains of TBS. One TBS train consists of 10 bursts of 4 stimulations at 100 Hz. The inter-burst interval is 200ms and the interval between trains is 2 s. fEPSPs responses were recorded for an hour after the stimulation protocol. All data were analyzed with an in-house program written with Matlab. Data are presented as mean ± SEM.

All animals were blinded and coded, and all recordings and analyses were conducted blindly. Animals were sedated by isoflurane inhalation and perfused intracardially with ice-cold choline chloride solution containing 124 mM choline chloride, 2.5 mM KCl, 26 mM NaHCO₃, 3.3 mM MgCl₂, 1.2 mM NaH₂PO₄, 10 mM Glucose, and 0.5 mM CaCl₂ (pH 7.4 equilibrated with 95% O₂ and 5% CO₂). Brains were then removed and placed in the same chilled choline chloride solution, and coronal acute slices of 300 µm from left and right hemispheres were collected and placed in oxygenated (95% O₂ and 5% CO₂) ACSF (in mM: NaCl 127, KCl 2.5, Glucose 10, NaHCO₃ 25, NaH₂PO₄ 1.25, MgCl₂ 2, CaCl₂ 2 mM) at 32°C for 1 hr and maintained at room temperature for the rest of the experiment. Whole-cell voltage clamp recordings of hippocampal neurons of Cre+ and Cre- Rab10 animals were performed with a Multiclamp 700B. Patch pipettes (3–6 ΩM) were filled with a Cs Methanesulfonate solution (in mM: Cs MeSO₄ 120, NaCl 5, TEA-Cl 10, HEPES 5, QX314-Br 5, EGTA 5, NaATP 4, MgGTP 0.3, pH 7.4). Experiments were performed at room temperature (18–20°C), and slices were perfused with oxygenated ACSF (in mM: NaCl 127, KCl 2.5, Glucose 10, NaHCO₃ 25, NaH₂PO₄ 1.25, MgCl₂ 2, CaCl₂ 2 mM, picrotoxin 100 μM). AMPAR and NMDAR-evoked responses were measured by voltage-clamping the cells at –70 mV and +40 mV, respectively. Postsynaptic currents were evoked by electrical stimulation using a concentric bipolar electrode (THC), with a pulse of 0.1ms. Input-output curves were first assessed by changing the stimulation intensity from 0 to 200 μA. For the rest of the recordings, the stimulation intensity was set to the amplitude that elicited a 50% EPSC response. Recordings were filtered at 2 kHz and digitized at 10 kHz. Series and input resistances were monitored throughout the experiment. All data were acquired and analyzed with an in-house program written in Matlab. Data are presented as mean ± SEM.

Results are reported as mean ± SEM. Statistical analysis was performed with GraphPad Prism 7 and 10. Comparisons between two groups were performed using unpaired two-tailed Student’s t-tests (* p<0.05, ** p<0.01, *** p<0.001, **** p<0.0001). Comparisons for more than two groups were performed using one-way ANOVA followed by Bonferroni’s multiple comparison tests or two-way ANOVA (* p<0.05, ** p<0.01, *** p<0.001, **** p<0.0001).