RanBP9 at the intersection between cofilin and Aβ pathologies: rescue of neurodegenerative changes by RanBP9 reduction

Molecular pathways underlying the neurotoxicity and production of amyloid β protein (Aβ) represent potentially promising therapeutic targets for Alzheimer's disease (AD). We recently found that overexpression of the scaffolding protein RanBP9 increases Aβ production in cell lines and in transgenic mice while promoting cofilin activation and mitochondrial dysfunction. Translocation of cofilin to mitochondria and induction of cofilin–actin pathology require the activation/dephosphorylation of cofilin by Slingshot homolog 1 (SSH1) and cysteine oxidation of cofilin. In this study, we found that endogenous RanBP9 positively regulates SSH1 levels and mediates Aβ-induced translocation of cofilin to mitochondria and induction of cofilin–actin pathology in cultured cells, primary neurons, and in vivo. Endogenous level of RanBP9 was also required for Aβ-induced collapse of growth cones in immature neurons (days in vitro 9 (DIV9)) and depletion of synaptic proteins in mature neurons (DIV21). In vivo, amyloid precursor protein (APP)/presenilin-1 (PS1) mice exhibited 3.5-fold increased RanBP9 levels, and RanBP9 reduction protected against cofilin–actin pathology, synaptic damage, gliosis, and Aβ accumulation associated with APP/PS1 mice. Brains slices derived from APP/PS1 mice showed significantly impaired long-term potentiation (LTP), and RanBP9 reduction significantly enhanced paired pulse facilitation and LTP, as well as partially rescued contextual memory deficits associated with APP/PS1 mice. Therefore, these results underscore the critical importance of endogenous RanBP9 not only in Aβ accumulation but also in mediating the neurotoxic actions of Aβ at the level of synaptic plasticity, mitochondria, and cofilin–actin pathology via control of the SSH1-cofilin pathway in vivo.

The defining pathological hallmark of Alzheimer's disease (AD) is the accumulation of amyloid β protein (Aβ) in brain associated with tau pathology, synapse loss, cytoskeletal aberrations, mitochondrial dysfunction, and cognitive decline. The generation of Aβ occurs via sequential βand γ-secretase processing of the amyloid precursor protein (APP) by beta site APP cleaving enzyme 1 (BACE1) and the presenilin (PS) complex, respectively. 1 Soluble oligomeric forms of Aβ are thought to be the most toxic species, resulting in synaptic loss and downstream neurotoxicity. 2 Despite the requirement for Tau in multiple aspects of Aβ-induced neurotoxicity, 3 a large knowledge gap exists as to how the Aβ oligomer-induced neurotoxic signals are transduced intracellularly to impair synaptic plasticity, eventually leading to neurodegeneration. Both Aβ and Tau promote cofilin-actin pathology, 4,5 cofilinactin pathology is widespread in AD brains, 6 and cofilin activity is also increased in AD brains. 7 Cofilin normally functions as a key regulator of actin dynamics that destabilizes filamentous actin (F-actin). Cofilin is inactivated by phosphorylation on Ser3 by LIM kinase 1 (LIMK1), whereas its dephosphorylation by Slingshot homolog 1 (SSH1) activates cofilin. 4 Upon oxidative stress and/or Ca 2+ elevation, 4,8,9 SSH1 is activated and active cofilin becomes oxidized on cysteine residues, resulting in rapid mitochondrial translocation to promote apoptosis and induction of cofilin-actin pathology. 10,11 An early and consistent impairment secondary to Aβ oligomer treatment in primary neurons is the shrinkage of dendritic spines 12 involving the rearrangement of F-actin cytoskeleton in spines and loss of spine-associated proteins such as postsynaptic density-95 (PSD95) and Drebrin, 13,14 as well as impaired mitochondrial function. 15, 16 We recently found that overexpression of the scaffolding protein RanBP9 increases Aβ production in cell lines and in transgenic mice. 17,18 Moreover, RanBP9 is significantly increased in brains of AD patients and the J20 APP transgenic model. 18,19 In studying the trafficking of APP, we also found that RanBP9 overexpression not only promotes the endocytosis of APP but also those of LRP and β1-integrin, the latter  resulting in disassembly of integrin-associated focal complexes (talin and vinculin). 20 In addition, RanBP9 overexpression promotes cofilin activation and the translocation of cofilin to mitochondria, resulting in overall mitochondrial dysfunction. 9,19 However, how RanBP9 activates cofilin is unknown, and it is not clear whether reduction in endogenous RanBP9 protects against Aβ oligomer-induced deficits in synaptic plasticity, cofilin-dependent pathology, Aβ accumulation, and memory impairment. Here we report that short interfering ribonucleic acid (siRNA) or genetic reduction in RanBP9 significantly reduces SSH1 levels and mitigates Aβinduced translocation of cofilin to mitochondria, cofilin-actin rod/aggregate formation, depletion of synaptic proteins, deficits in synaptic plasticity, Aβ accumulation, and contextual memory deficits in vivo.

Results
Endogenous RanBP9 mediates Aβ42 oligomer-induced translocation of cofilin to mitochondria and promotes cofilin activation via positively regulating SSH1. We assessed whether Aβ oligomers alter cofilin translocation to mitochondria and whether siRNA knockdown of RanBP9 may influence this phenotype. We prepared Aβ1-42 oligomers precisely as previously characterized, 21 Figure  S1A). In vivo, RanBP9 levels were greatly decreased in the hippocampus of 3-month-old APP/PS1;RanBP9+/ − mice in both cytosol and mitochondrial fractions compared with APP/PS1 mice 22 (Figure 1d). Although cofilin levels were unchanged in the cytosol fraction between the genotypes, cofilin was significantly reduced in the hippocampal mitochondrial fraction of APP/PS1;RanBP9+/ − mice compared with APP/PS1 mice (Figures 1d and e), indicating that endogenous RanBP9 facilitates translocation of cofilin to mitochondria in vivo. As expected, 3-month-old RanBP9+/ − mice exhibited reduced RanBP9 protein together with significantly increased inactive phospho-cofilin without altering total cofilin (Figures 1f and g), which was accompanied by significantly decreased SSH1 protein in RanBP9+/ − mice (Figures 1f and h), indicating that RanBP9 positively regulates SSH1 levels. Indeed, SSH1 but not LIMK1 levels were significantly increased secondary to RanBP9 overexpression in HT22 cells ( Figures 1I and j). Consistent with these observations, RanBP9 not only markedly increased SSH1 but also co-immunoprecipitated with SSH1 in brain ( Figure 1k). Accordingly, siRNA knockdown of RanBP9 in HT22 cells strongly reduced the enhancement of cofilin-SSH1 complex induced by Aβ42 O treatment (Figure 1l), while significantly decreasing SSH1 levels ( Figures 1I and m). In days in vitro 21 (DIV21) primary cortical neurons, Aβ42 O (2 h) markedly increased RanBP9-SSH1 and cofilin-SSH1 complexes (Figure 1n), suggesting that RanBP9 not only increases SSH1 levels but may also serve to facilitate cofilin-SSH1 interaction upon Aβ42 O exposure. To determine whether the RanBP9-SSH1 complex alters SSH1 protein stability, we performed cycloheximide turnover experiments. Indeed, RanBP9 siRNA and overexpression markedly accelerated and delayed the turnover of SSH1, respectively ( Figure 1o), confirming that RanBP9 positively regulates SSH1 protein stability.
Endogenous RanBP9 and SSH1 promote cofilin-actin rod formation and Aβ42 oligomer-induced collapse of growth cones in primary hippocampal neurons. Hyperactive cofilin can form cofilin-saturated actin filament bundles (or rods) when challenged by oxidative stress, excitotoxic insult, or Aβ oligomers in primary neurons. These cofilin-actin rods accumulate in neurites, which blocks neuritic transport of key cargo to distal locations. 23,24 To assess the role of RanBP9 in cofilin-actin rod formation, we transiently transfected green fluorescent protein (GFP)-cofilin in wild-type (WT) and RanBP9+/ − hippocampal neurons. On DIV8, we treated neurons with a low concentration of hydrogen peroxide (1 μM) for 30 min, which is known to induce cofilinactin rod formation. WT neurons formed significantly higher numbers of GFP-cofilin rods per neuron and per neuronal cytoplasmic area (NCA) compared with RanBP9+/ − neurons (Figures 2a and b). We next treated DIV8 WT and RanBP9+/ − neurons with or without Aβ42 O (1 μM) for 24 h, which has been shown to induce cofilin-actin rod formation in 10-20% of neurons. 25  Endogenous RanBP9 mediates the depletion of postsynaptic proteins induced by Aβ42 oligomers in primary hippocampal neurons. Dendritic spines are structures critical for excitatory synaptic transmission and highly enriched in postsynaptic proteins such as Drebrin, PSD95, and F-actin. 26 To assess synaptic perturbations induced by   Significant increase in endogenous RanBP9 in APP/PS1 mice brains. We examined endogenous RanBP9 expression in the APP/PS1 mice expressing the 'Swedish' and PS1ΔE9 mutation by immunoblotting and immunohistochemistry. Endogenous RanBP9 protein levels were significantly increased by 3.5-fold in the hippocampus of 8-month-old APP/PS1 mice by immunoblotting (Figures 4a  and b), consistent with our previous observations in 12-month-old J20 mice harboring the 'Swedish' mutation and in AD brains. 18,19 Further, immunohistochemical analysis of APP/PS1 mouse hippocampus demonstrated greatly enhanced RanBP9 levels in the dentate gyrus (DG) and CA3 regions compared with WT littermates (Figure 4c).
Rescue of neuroinflammation, synaptic damage, and Aβ accumulation by RanBP9 reduction in APP/PS1 mice. Given the robust upregulation of RanBP9 levels in APP/PS1 mice, we next determined whether genetic reduction in RanBP9 can prevent neuroinflammation and synapse loss associated with APP/PS1 transgenic mice. We performed immunohistochemistry for glial fibrillary acidic protein (GFAP) and Iba1 to detect activated astrocytes and microglia as well as Synapsin I and PSD95 to detect pre-and postsynaptic proteins from 8-month-old APP/PS1, APP/PS1;RanBP9+/ − , and WT littermates. As expected, APP/PS1 mice demonstrated significantly increased GFAP and Iba1 immunoreactivities throughout the hippocampus and areas of the anterior cortex associated with Aβ accumulation as well as diminished PSD95 and Synapsin I immunoreactivities within the stratum lucidium (SL: synaptic terminating zone) of CA3 (Figures 5a  and b). In contrast, APP/PS1;RanBP9+/ − mice showed significantly reduced GFAP and Iba1 immunoreactivities in the hippocampus and anterior cortex, as well as significantly rescued Synapsin I and PSD95 immunoreactivities within the SL of CA3 compared with APP/PS1 littermate mice, essentially indistinguishable from WT mice (Figures 5a and b). As RanBP9 overexpression promotes Aβ production in cultured cells and in vivo, 17,18 we next examined whether endogenous RanBP9 reduction alters Aβ burden in the hippocampus and anterior cortex of 8-month-old APP/PS1 mice. Indeed, APP/PS1;RanBP9+/ − mice demonstrated a significant decrease in both the area covered by total Aβ deposits and thioflavin S-positive fibrillar Aβ deposits within the hippocampus and anterior cortex compared with  (Figures 6a and c). In addition, soluble Aβ levels were also significantly reduced in the hippocampus of APP/PS1;RanBP9+/ − mice compared with littermate APP/PS1 mice (Figures 6d and e). These results show that RanBP9 reduction protects against Aβ accumulation, as well as neuroinflammation and synaptotoxicity in vivo.
Accumulation of cofilin-actin rods in APP/PS1 mice and significant attenuation of cofilin-actin rod formation by RanBP9 reduction. Given the positive regulation of SSH1 and cofilin, as well as cofilin-actin rods in primary neurons by RanBP9, we assessed cofilin-actin rods/aggregates in WT, APP/PS1, and APP/PS1;RanBP9+/ − brains. For this, we used Sudan Black B to quench background and the more diffuse non-rod staining. Therefore, only intense signals, such as rods, are visualized, while removing normal cytoplasmic immunoreactivity. Using this method, we found that cofilin rods/aggregates were readily detectable in the hippocampus and anterior cortex of 9-month-old APP/PS1 mice, while very few were found in WT mice (Figure 7a). Such cofilin rod/ aggregate structures, which colocalized with actin, were significantly increased by~10-fold in APP/PS1 versus WT mice (Figures 7a and c) and resembled those seen in AD brains (Figure 7d). RanBP9 reduction significantly attenuated cofilin-actin rod/aggregate formation, although not completely to the level of WT mice (Figures 7a and c). In the AD entorhinal cortex, cofilin rod/aggregate structures were numerous and largely did not colocalize with 12E8-positive phospho-Tau immunoreactivity, albeit within the same region and in close proximity (Figure 7d). These findings in vivo confirm our findings in primary neurons that RanBP9 reduction ameliorates cofilin-actin pathology in an in vivo mouse model of Aβ accumulation. RanBP9 reduction rescues deficits in synaptic plasticity and contextual memory associated with APP/PS1 mice. We tested short-term and long-term synaptic plasticity from acute hippocampal slices prepared from 3-month-old WT, APP/PS1, RanBP9+/ − , and APP/PS1;RanBP9+/ − mice, an age when little to no Aβ plaques are detected. 22 The stimulating electrode was placed in the Schaffer collaterals of the hippocampus, and the recording electrode was positioned at the CA1 stratum radiatum below the pyramidal cell layer. As shown in Figure 8a, input-output analysis did not markedly differ among WT, APP/PS1, and APP/PS1; RanBP9+/ − , and RanBP9+/ − slices. However, the APP/ PS1;RanBP9+/ − slices showed a nonsignificant decrease in basal synaptic transmission compared with WT, RanBP9+/ − , and APP/PS1 slices (Figure 8a). In paired pulse facilitation (PPF) experiments, significant differences were observed in fEPSP slope across genotypes among all interstimulus intervals (Figure 8b). Correction for multiple comparisons surprisingly showed that APP/PS1;RanBP9+/ − slices exhibited significantly higher PPF compared with both WT, RanBP9+/ − , and APP/PS1 slices across most interstimulus intervals (Figure 8b), indicating a synergistic effect of RanBP9 and APP/PS1 genotypes on cooperative presynaptic efficacy. For long-term potentiation (LTP) measurements, we detected no differences in fEPSP slope among WT, APP/PS1, RanBP9+/ − , and APP/PS1;RanBP9+/ − slices at baseline (Figure 8c). However, after theta burst stimulation, we observed significant differences in fEPSP slope across genotypes for all time points (Figure 8c). Correction for multiple comparisons showed that APP/PS1 slices were significantly impaired in fEPSP slope compared with WT, RanBP9+/ − , and APP/PS1; Similarly, RanBP9+/ − slices exhibited significantly stronger LTP than WT slices up to 30 min and did not significantly differ from APP/PS1;RanBP9+/ − slices (Figure 8c). Therefore, these results indicate that RanBP9 reduction not only rescues the deficits in synaptic plasticity associated with the APP/PS1 mice but also further potentiates synaptic plasticity.
To determine whether the changes in LTP and synaptic proteins correlate with learning and memory, we carried out fear conditioning tests (contextual and cued), in which WT, APP/PS1, and APP/PS1;RanBP9+/ − mice were trained on day 1 for both. On day 2, we tested mice for contextual and cued conditioning memory. As shown in Figure 8d, we observed significant differences across genotypes in the amount of time spent freezing (in seconds) in the contextual fear conditioning test (Figure 8d). Specifically, WT mice froze for significantly longer than APP/PS1 mice but did not significantly differ from the APP/PS1;RanBP9+/ − mice (Figure 8d), as the latter mice showed an intermediate amount of time freezing. However, we did not observe significant differences across genotypes in the hippocampus-independent cued conditioning freezing times (Figure 8e). We also did not find significant differences across the three genotypes in open field or rotarod tests (Figures 8f and g), indicating the these genotypes do not alter general locomoter activity or coordination. Taken together, these results show that a threshold of endogenous RanBP9 is required to mediate the full deficits in hippocampus-dependent contextual memory but not cued memory.

Discussion
Molecular pathways that govern the neurotoxicity and production of Aβ are attractive therapeutic targets for AD. In this study, we utilized HT22 cells, primary neurons, acute brain slices, and genetically modified mice to gain insights into the role of endogenous RanBP9 in Aβ-induced synaptic degeneration, cofilin activation, cofilin-actin rod formation, and Aβ accumulation in a mouse model of AD. We made a number of novel observations demonstrating that endogenous RanBP9 positively regulates SSH1 levels and is critical for Aβ-induced translocation of cofilin to mitochondria, cofilin-actin rod/aggregate formation, collapse of growth cones, loss of synaptic proteins, and impairments in synaptic plasticity and learning/memory. Our findings underscore the critical importance of the RanBP9-SSH1-cofilin pathway in AD pathogenesis. A prerequisite for cofilin translocation to mitochondria and formation of cofilin-actin rods is its activation via dephosphorylation and oxidation on several cysteine residues. 10,11 Therefore, the observation that Aβ oligomers induced cofilin translocation to mitochondria and formation of cofilin-actin rods infers its rapid activation and oxidation by Aβ oligomers. Interestingly, Aβ O also increased RanBP9 translocation to mitochondria, which may in part be explained by its association with the pro-apoptotic mitochondria-associated protein P73. 27 Genetic reduction and siRNA knockdown of RanBP9 significantly mitigated both of these measures and reduced cofilin activation. As cofilin is activated by SSH1-dependent dephosphorylation and inactivated by LIMK-dependent phosphorylation, it is plausible that RanBP9 alters one or both of these pathways. Indeed, we found that RanBP9 positively regulates SSH1 but not LIMK1 levels in HT22 cells and in brain, which accounts for increased cofilin activation by RanBP9. As RanBP9 levels are robustly increased in APP/ PS1 mouse brains, it is likely that Aβ O also activates cofilin via a mechanism involving RanBP9 and SSH1. Aβ is known to increase cytosolic Ca 2+ and activate calcineurin as well as enhance production of reactive oxygen species (ROS), 28,29 both of which are known activators of SSH1. Calcineurindependent dephosphorylation of SSH1 serves to activate SSH1, 30 and cysteine oxidation of 14-3-3ζ/τ removes the inhibition of SSH1 by 14-3-3ζ/τ. 8 In addition, cysteine oxidation of cofilin itself is required for both mitochondrial translocation and cofilin-actin rod formation. 10,11 We recently showed that Aβ O promotes cofilin-actin rod formation via a pathway requiring PrP c and NADPH oxidase (NOX), the latter which is involved in the generation of ROS. 31 We have also shown that Aβ O and RanBP9 impair mitochondrial Ca 2+ buffering and promote mitochondrial superoxide production in a cofilindependent manner. 9,19 Therefore, the engagement of Aβ O to its neuronal receptors likely involves downstream Ca 2+ and ROS from multiple sources. Activated/oxidized cofilin likely contributes to sustaining this cycle by promoting mitochondrial superoxide production. We observed that RanBP9+/ − neurons are resistant to Aβ oligomer-induced collapse of F-actin containing growth cones in immature DIV9 neurons and depletion of postsynaptic proteins (Drebrin, PSD95, and F-actin) in mature DIV21 neurons. Such findings are likely explained, at least in part, by the reduction in cofilin activation status in RanBP9+/ − neurons. Specifically, phosphorylation and dephosphorylation of cofilin are associated with dendritic spine growth and shrinkage during LTP and LTD, respectively, [32][33][34] and cofilin regulates the trafficking of α-amino-3-hydroxy-5-methyl-4isoxazolepropionic acid (AMPA) receptors in dendritic spines. 35 Further, β-arrestin 2 recruits cofilin to dendritic spines upon N-methyl-D-aspartate receptor activation to control the remodeling of spines, and β-arrestin 2 knockout neurons are resistant to Aβ-induced dendritic spine loss through spatial control of cofilin activation. 36 Despite the crucial involvement of cofilin in synaptic plasticity, the aforementioned mechanism alone is unlikely to explain the full extent of the rescue and enhancement of LTP and PPF associated with APP/PS1;RanBP9+/ − mice, as both LTP and PPF in were strongly enhanced significantly beyond those of WT slices. However, in the absence of the APP/PS1 genotype, RanBP9+/ − mice exhibited normal PPF but significantly enhanced LTP. The exaggerated PPF in APP/PS1;RanBP9+/ − slices may be explained by lower probability of Ca 2+ -evoked neurotransmitter release as evidenced by somewhat reduced fEPSP slope per stimulus amplitude (input/output curve). However, the second paired Ca 2+ signal appears to overcompensate to release a larger pool of neurotransmitter. We hypothesize that RanBP9 reduction enhances synaptic associativity and cooperativity perhaps at the level of synapse formation and maintenance. Indeed, RanBP9+/ − slices produced significantly enhanced LTP compared with WT and did not differ significantly from APP/PS1;RanBP9+/ − slices. Overexpression of RanBP9 significantly reduces axon elongation in the chick spinal cord via Plexin-A, 37 and we also showed in this study that reduction in RanBP9 increases neurite arborization/complexity in mature neurons and protects against both Aβ oligomer-induced growth cone collapse and dendritic postsynaptic protein depletion. Previous studies have also shown the involvement of RanBP9 in negatively regulating β1-integrin function 20 and a role in the nucleocytoplasmic regulation of cell morphology via its interaction with muskelin. 38 Therefore, RanBP9 may be involved in regulating neuritogenesis, synaptic maintenance, and synaptic plasticity changes via these pathways in addition to cofilin activation.
RanBP9 reduction protected against both Aβ O -and hydrogen peroxide-induced cofilin-actin rod formation in primary neurons, thereby indicating a direct effect of RanBP9 and SSH1 reduction on cofilin-actin rods. Indeed, SSH1 reduction per se significantly mitigated Aβ42 O -induced cofilin rod/ aggregate formation and collapse of growth cones, indicating that SSH1 reduction, at least in part, is responsible for such effects of RanBP9. In vivo, however, RanBP9 reduction mitigated both cofilin-actin rod/aggreagate formation and Aβ accumulation/plaques. Therefore, it is likely that RanBP9 reduction lowers cofilin-actin rod/aggregate formation both directly via SSH1 regulation and indirectly by reducing Aβ accumulation. Cofilin-actin rods are thought to be an initial protective response to oxidative insults; 39 however, persistent accumulation of rods physically blocks neuritic transport of cargo and impairs synaptic activity. 23,24 Inhibition of RanBP9 and SSH1 may not be sufficient to fully block cofilin-actin pathology, as ROS pathways acting directly on cofilin may also promote cofilin-actin pathology. In this light, it is intriguing that Tau overexpression induces cofilin-actin pathology and potently increases F-actin bundling, 5 while disrupting mitochondrial fission via an actin-dependent mechanism. 40 Taken together, this study demonstrated the crucial involvement of endogenous RanBP9 in Aβ-induced cofilin activation via SSH1 and downstream consequences on cofilin-actin rod formation, Aβ accumulation, synaptic plasticity, and contextual memory. These findings underscore the role of RanBP9 and its associated molecular pathway as critical mediators of AD pathology and potential therapeutic targets.

Materials and Methods
Cells, complementary deoxyribonucleic acid constructs, siRNA sequences, and transfections. Hippocampus-derived HT22 cells were maintained in Dulbecco's modified Eagle's medium containing 10% FBS. HT22 cells were obtained from Professor David Schubert (Salk Institute, San Diego, CA, USA). The RNAi targeting RanBP9 (5′-UCUUAUCAACAAUACCUGC-3′) and SSH1 (5′-GAGGAGCUGUCCCGAUGAC-3′) were obtained from GE Dharmacon (Lafayette, CO, USA). All cells were transfected for 48 h using lipofectamine 2000 (Invitrogen, Carlsbad, CA, USA) according to the manufacturer's instructions with siRNA duplexes or control single-stranded denatured control siRNAs before performing biochemical and/or immunocytochemical assays. All siRNAs were transfected at a final concentration of 100 nM. Cortical and hippocampal primary neurons were derived from postnatal day 0 (P0) pups and grown on poly-D-lysine-coated coverslips or plates as previously described. 19 Quantitation of digitized immunoblotting data was performed using the Image J software (NIH Image J, Bethesda, MD, USA). , and fluorescently labeled secondary antibodies (Invitrogen) were obtained from the indicated sources. The mouse monoclonal anti-RanBP9 antibody was a generous gift from Professor Elizabetta Bianchi (Pasteur Institute, France). Synthetic Aβ1-42 peptide was purchased from American Peptide (Sunnyvale, CA, USA). Aβ1-42 oligomers were prepared as previously characterized. 21 Briefly, Aβ1-42 powder was dissolved in hexafluoro-2-propanol (HFIP) at 1 mM for 30 min at room temperature, aliquoted to eppendorf tubes, allowed to evaporate overnight in fume hood, and subjected to speed vacuum for 1 h to remove traces of HFIP or moisture. To prepare Aβ oligomers, Aβ1-42 film was then dissolved in dimethyl sulfoxide (5 mM), and F-12 cell culture medium (without phenol) was added to a final concentration of 100 μM Aβ1-42 and incubated at 4°C for 24 h.
Cell/tissue lysis and immunoblotting. Cultured cells or brain homogenates were lysed with lysis buffer (50 mM Tris-Cl, 150 mM NaCl, 2 mM ethlenediaminetetraacetic acid, and 1% TritonX-100). For isolation of mitochondria, mitochondrial isolation kit (Thermo Scientific, Rockford, IL, USA) was used according to the manufacturer's instructions. Protein quantification was performed by a colorimetric detection reagent (BCA Protein Assay, Pierce, Rockford, IL, USA). Equal amounts of protein were subjected to SDS-polyacrylamide gel electrophoresis (PAGE) and transferred to nitrocellulose membranes for immunoblotting. After probing with the primary antibody, the corresponding peroxidase-conjugated secondary antibody was detected by electrochemiluminescence (ECL) western blot reagents (Pierce).
Mice. APP/PS1, WT, and RanBP9+/ − mice 19 were all bred in the C57BL6 background for at least three generations before interbreeding with each other. APP/PS1 mice, which express 'Swedish' APP and PS1 ΔE9 mutations, were obtained from Jackson Immunoresearch (West Grove, PA, USA). 22 Immunofluorescence. Immunohistochemistry and immunocytochemistry were performed as previously described. 19 Briefly, animals were perfused with 4% paraformaldehyde in PBS, and brains were post-fixed in the same fixative for 24 h. The brains were then cryoprotected in 30% sucrose and sectioned (30 μm) on a cryostat or microtome. For primary neurons and HT22 cells, cells were fixed in 4% paraformaldehyde for 15 min at room temperature. After blocking with normal goat serum, primary antibodies were applied overnight at 4°C, and secondary antibodies were applied for 45 min at room temperature, followed by counterstaining with Hoechst33342 or DAPI and mounting. For detection of cofilin-actin rods in brain, 70% ethanol was applied to sections for 5 min after washing the secondary antibody, followed by 0.1% Sudan Black B in 70% ethanol for 12 min, two washes with 70% ethanol, several washes in Tris-buffered saline (TBS), and rinsed in H 2 0 before mounting. All immunoreactivities were quantitated from every 12th serial section through an entire hippocampus or anterior cortex. All images were acquired with the Olympus FV10i confocal microscope (Tokyo, Japan) and quantitated using the Image J software. Comparison images were acquired with identical laser intensity, exposure time, and filter. Adjustments to the brightness/contrast were applied equally to all comparison images.
Behavioral analysis. Fear conditioning (contextual and cued), rotarod, and open field tasks were performed as previously described. 41 For fear conditioning, an aversive stimulus (in this case a mild foot shock, 0.5 mA) was paired with an auditory conditioned stimulus (white noise) within a novel environment. Training consisted of two mild shocks paired with two conditioned stimuli with a 3-min interval between each shock. Freezing on the training day in response to the foot shock was used as an estimate of learning during the acquisition trial. To test conditioning to the context, animals were re-introduced to the same training chamber for 6 min and freezing behavior was recording by tracking software (Any-maze, Wood Dale, IL, USA) every second. To test conditioning to the tone, animals were introduced to a novel context, consisting of a chamber with different shape, floor and olfactory cues from the training chamber. Mice were scored for 3 min, before and after the tone in the same manner described above. Learning was assessed by measuring freezing behavior (i.e., motionless position) every second. The open field was used as a standard test of general activity. Briefly, animals were monitored for 15 min in a 40 cm square open field with a video tracking software (Any-maze). General activity levels were evaluated by measurements of total distance traveled. Motor performance was evaluated by an accelerating rotarod apparatus with a 3 cm diameter rod starting at an initial rotation of 4 r.p.m. slowly accelerating to 40 r.p.m. over 5 min. The time spent on the rod during each of four trials per day for 2 consecutive days was measured previously described in. 42 Electrophysiology. Hippocampus slices were prepared from 3-month-old WT, APP/PS1, and APP/PS1;RanBP9+/ − mice and subjected to input/output curved, paired pulse facilitation, and LTP as previously described. 43 Briefly, animals were killed, brains were harvested, and sectioned horizontally (400 μm) in ice-cold cutting solution (110 mM sucrose, 6 mM NaCl, 3 mM KCl, 26 mM NaHCO 3 , 1.25 mM NaH 2 PO 4 , 7 mM MgCl 2 , 0.5 mM CaCl 2 , 10 g/l glucose, pH 7.3-7.4). The hippocampus was dissected and acclimated in 50:50 solution (cutting:artificial cerebrospinal fluid (ACSF)) for 10 min at room temperature. Then the slices were transferred to ACSF (125 mM NaCl, 2.5 mM KCl, 1.25 mM NaH 2 PO 4 , 0.26 mM NaHCO 3 , 1.2 mM MgCl 2 , 2.0 mM CaCl 2 , and 10 g/l glucose, pH 7.3-7.4, saturated with 95% O 2 and 5% CO 2 ). Slices were recovered in ACSF at room temperature at least 40 min, followed by a final incubation in ACSF for 1 h at 30°C. Extracellular field potential recording, LTP: the recording chamber was held at 30 ± 0.5°C with ACSF flow rate of 1 ml/min. The stimulating electrode was placed in the Schaffer collaterals of the hippocampus. The recording glass electrode loaded with ACSF was positioned at the CA1 stratum radiatum below the pyramidal cell layer. Stimulating pulses were generated by the Digidata 1322A interface (Molecular Devices, Sunnyvale, CA, USA) and a stimulus isolator (model 2200; A-M Systems, Sequim, WA, USA) under control of Clampex 10.0 software (Molecular Devices). Field excitatory postsynaptic potentials (fEPSP) were amplified using a differential amplifier (model 1800; A-M Systems), filtered at 1 kHz, and digitized at 10 kHz.
Input-output analysis was performed by stepping stimulation amplitude from 1 to 15mV. Stimulation amplitude that elicited half-maximal fEPSP was determined by the input-output curve and stimulation rate of 0.05 Hz was used through the whole experiment. PPF, which is short-term plasticity, was evoked by two pulses with interpulse intervals from 20 to 300 ms. Percentage of the facilitation was calculated by dividing fEPSP slope elicited by the second pulse with the fEPSP slope elicited by the first pulse. LTP was induced by theta burst stimulation (TBS) (five trains of four pulses at 200 Hz separated by 200 ms, repeated six times with an inter-train interval of 10 s). LTP was sampled 60 min after the induction, and calculated by dividing the slope of 60 min post-induction responses with the average slope of 20-min baseline responses.