Wnt5a Is Essential for Hippocampal Synaptic Plasticity.

The dramatic changes in adult neuronal morphology with Wnt5a loss prompted us to ask whether Wnt5a is essential for hippocampal synaptic transmission in vivo. We performed electrophysiological recordings in CaMKII-Wnt5a^fl/fl mice at 3 mo of age, a time when CA1 neuronal morphology is still intact, and at 6 mo when the morphology is impaired. We observed that basal synaptic transmission is normal in CaMKII-Wnt5a^fl/fl mice at 3 mo but not at 6 mo, consistent with impaired dendritic structures at this age (Fig. 2 A–D). We also measured the probability of synaptic release at presynaptic sites by paired pulse facilitation (PPF) analyses and found that PPF was comparable between CaMKII-Wnt5a^fl/fl mice and control littermates at both 3 and 6 mo of age (Fig. S3). The normal presynaptic properties in Wnt5a-deficient mice suggests that Wnt5a acts primarily at postsynaptic sites.

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Fig. 2.

Wnt5a loss impairs LTP in the absence of changes in basal synaptic transmission or neuronal morphology. (A–D) Basal synaptic transmission is unaffected in CaMKII-Wnt5a^fl/fl (KO) mice at 3 mo but impaired at 6 mo compared with Wnt5a^fl/fl litter-mates (WT). The input–output relationships and slopes of input–output curves are normal in 3-mo-old KO mice, but significantly different from WT at 6 mo. Results are mean ± SEM from five animals per genotype (n = 18 slices for WT and 17 slices for KO mice) at 3 mo, and from five WT (n = 21 slices) and four KO mice (n = 23 slices) at 6 mo. **P < 0.01, two-tailed t test. [Scale bars, 1 mV (vertical), 2.5 ms (horizontal) for all sample traces.] (E) Impaired LTP at the Schaffer collateral–CA1 synapses in 3-mo-old KO mice. n = 11 slices from 5 WT mice and n = 13 slices from 6 KO mice. Sample traces represent fEPSPs (field excitatory postsynaptic potentials) right before (dashed line) and 1 h after (solid line) θ-burst stimulation. [Scale bars, 1 mV (vertical), 10 ms (horizontal) for both sample traces.] (F and G) Induction and maintenance of LTP are attenuated in KO mice. Results are mean ± SEM from five WT mice (n = 11 slices) and six KO mice (n = 13 slices). *P < 0.05, ***P < 0.001, two-tailed t test. (H) Low-frequency stimulation-induced LTD at Schaffer collateral–CA1 synapses is unaltered in 3-mo CaMKII-Wnt5a^fl/fl mice. Sample traces represent fEPSPs right after (dashed line) and 1hr after (solid line) stimulation. [Scale bars, 1 mV (vertical), 10 ms (horizontal) for both sample traces.] (I) Mean fEPSP slopes are comparable between the genotypes. Results are mean ± SEM from five Wnt5a^fl/fl mice (n = 14 slices) and five CaMKII-Wnt5a^fl/fl mice (n = 16 slices).

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Fig. S3.

Wnt5a deletion does not affect presynaptic properties. (A and B) Presynaptic vesicle release is normal in 3- and 6-mo-old CaMKII-Wnt5a^fl/fl (KO) mice as measured by paired-pulse ratios with different interstimulus interval (ms). Results are means ± SEM from four animals per genotype for 3-mo-old animals [n = 17 slices for Wnt5a^fl/fl (WT) and n = 15 slices from KO mice]. For 6-mo-old animals, four WT mice (n = 24 slices) and three KO mice (n = 22 slices) were used. [Scale bars, sample traces in Insets; 1 mV (vertical), 10 ms (horizontal).] Statistical analyses were done using a two-way ANOVA.

Hippocampal neurons exhibit prominent synaptic plasticity in which activity-dependent modulation of the strength of synaptic connections underlies learning and memory (36). Previously, broad-spectrum Wnt antagonists have been shown to affect synaptic structure, plasticity, and cognitive functions in adult organisms (37–41). However, there are 19 vertebrate Wnts, and which Wnt is essential for these functions in the adult brain in vivo remains unknown. To assess the role of Wnt5a in NMDA receptor-dependent long-term potentiation (LTP), an electrophysiological correlate of strengthening of synaptic transmission, we used θ-burst stimulation to induce LTP at Schaffer collateral–CA1 synapses in 3-mo-old mice. Recordings from Wnt5a^fl/fl control slices revealed a robust induction of LTP and a sustained maintenance phase (Fig. 2 E–G). In contrast, CaMKII-Wnt5a^fl/fl slices showed a significant reduction in both induction (274.3 ± 18.2% in control slices vs.. 201 ± 7.9% in CaMKII-Wnt5a^fl/fl slices, P = 0.0002) and maintenance (190.8 ± 13.3% in control slices vs. 151.9 ± 5.4% mutant slices, P = 0.01) phases of LTP (Fig. 2 E–G). The impairment in LTP detected at 3 mo in CaMKII-Wnt5a^fl/fl mice when neuronal structure and basal synaptic transmission are still intact, suggests that synaptic plasticity is more susceptible to the loss of Wnt5a.

To address if Wnt5a contributes to NMDA receptor-dependent long-term depression (LTD), an electrophysiological correlate of weakening of synaptic transmission, we used a standard low-frequency stimulation paradigm to induce LTD in the CA1 hippocampus. In contrast to the LTP defect, we found no differences in LTD between mutant and control mice at 3 mo of age (Fig. 2 H and I). Taken together, these results indicate a specific role for Wnt5a in the potentiation of synaptic efficacy.

Wnt5a Is Essential for Spatial Learning and Memory.

Synaptic plasticity is widely considered to be a cellular mechanism that underlies learning and memory (42, 43). In addition, structural maintenance of synaptic connectivity has been postulated to be critical for life-long memories (44). Given the decreased CA1 LTP in 3-mo-old CaMKII-Wnt5a^fl/fl mice, we subjected Wnt5a mutant mice to behavioral paradigms to evaluate cognitive functions. In the novel-object recognition test, which evaluates the preference of mice to explore a new over a familiar object (45), adult Wnt5a^fl/fl mice showed a significant preference for the novel object, with 3- and 6-mo-old mice spending 70.8 ± 5.5% and 68.1 ± 8.1% of time exploring the novel object, respectively. However, CaMKII-Wnt5a^fl/fl mice showed no preference for the novel object at both 3 and 6 mo of age (Fig. 3 A and B), indicating a deficit in recognition memory.

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Fig. 3.

Adult Wnt5a mutant mice show a progressive decline in hippocampus-mediated behaviors. (A and B) WT mice show a significant preference for exploring a new object in the novel-object recognition task, whereas Wnt5a KO mice spent similar amounts of time with familiar and new objects. Deficits in recognition memory were evident in both 3- and 6-mo-old mutants. Dashed line indicates equal amount of time spent exploring new and familiar objects. Results are mean ± SEM from n = 21 WT mice and n = 19 KO mice at 3 mo, and n = 16 WT mice and n = 23 KO mice at 6 mo. *P < 0.05, **P < 0.01 significantly different from 50% time spent with the novel-object, two-tailed t test. (C) Timeline for the Morris water maze (MWM) tasks. (D) Three-month-old KO mice show impaired spatial learning in the Morris water maze test. Results are mean ± SEM from n = 20 WT and n = 19 KO mice. *P < 0.05, **P < 0.01, two-way ANOVA with Fisher’s least-significant difference (LSD) post hoc test. (E) Six-month-old KO mice exhibit more severe deficits in learning and fail to acquire the latency of control animals even on day 12. Results are mean ± SEM from n = 18 WT and n = 20 KO mice. *P < 0.05, **P < 0.01, ***P < 0.001, two-way ANOVA with Fisher’s LSD post hoc test. (F and G) During probe trials, 3- and 6-mo-old KO mice showed less preference for the target quadrant compared with WT mice. Results are mean ± SEM, *P < 0.05, **P < 0.01, two-tailed t test. (H and I) During the memory retention test, both 3- and 6-mo-old KO mice spent less time in the target quadrant compared with control littermates. However, older mutant mice exhibited a marked decay in memory retrieval by the fifth day of the probe trial. White dashed line indicates 25% of time that mice spent in the target quadrant. Results are mean ± SEM; *P < 0.05, **P < 0.01, ***P < 0.001 significantly different from control littermates; ^#P < 0.05 significantly different from 25% line, two-tailed t test.

To test hippocampus-dependent spatial learning in these mice, we used the Morris water maze to test an animal’s ability to use spatial cues to locate a hidden platform in a tank of water (46) (Fig. S4A). Importantly, CaMKII-Wnt5a^fl/fl mice had visual acuity and swimming speeds comparable to littermate controls (Fig. S4 B and C). Three-month-old CaMKII-Wnt5a^fl/fl mice took significantly longer time to locate the hidden platform during the initial training period of 12 consecutive days using four trials per day (see schematic in Fig. 3C), compared with age-matched controls, eventually achieving similar latencies as control mice on the sixth day (Fig. 3D). Six-month-old CaMKII-Wnt5a^fl/fl mice also required significantly more time to find the platform. However, 6-mo-old mutants did not achieve similar latencies as control mice, even when tested on the 12th day (Fig. 3E). These results show that acquisition of spatial learning is impaired in the absence of structural deficits in 3-mo-old Wnt5a mutant mice, but that learning deficits are exacerbated with the appearance of morphological abnormalities in older animals.

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Fig. S4.

Behavioral deficits in adult Wnt5a mutant mice. (A) Schematic of the Morris water maze. (B and C) Swimming speeds and visual acuities are similar between CaMKII-Wnt5a^fl/fl mice (KO) and control Wnt5a^fl/fl litter-mates (WT) at both 3 and 6 mo of age. Results are means ± SEM from 20 WT, 19 KO mice at 3 mo of age, and 17 WT, 20 KO mice at 6 mo. (D and E) Three- and 6-mo-old KO mice exhibit fewer crossings over the original platform location during the probe trial compared with control littermates. Results are means ± SEM from n = 20 WT and n = 19 KO mice at 3 mo, and from n = 18 WT and n = 20 KO mice at 6 mo. *P < 0.05, two-tailed t test. (F and G) Both 3- and 6-mo-old KO mice showed deficits in reversal learning compared with WT mice, although the deficits were more pronounced in older mutant animals. *P < 0.05, **P < 0.01, ***P < 0.001, two-way ANOVA with Fisher’s LSD post hoc test.

To evaluate reference memory, we conducted a probe trial on day 13 in which the platform was removed and measured the amount of time that mice spent in the original target quadrant (Fig. 3C). Both 3- and 6-mo-old mutant mice spent significantly less time in the target quadrant and made fewer platform crossings (Fig. 3 F and G and Fig. S4 D and E). We then performed a reversal training by relocating the hidden platform to the opposite quadrant from days 14–25 (Fig. 3C). As in the intial training phase, 3-mo-old Wnt5a mutant mice required more days of training to find the new platform, but eventually reached similar latencies as control mice (Fig. S4F). However, 6-mo-old mutant mice took significantly longer to find the platform compared with control littermates even on day 25 (Fig. S4G). We then conducted probe trials for 7 d (days 26–32) to assess memory retention. Three-month-old mutants maintained a preference for the target quadrant for the 7 d of the probe trial (Fig. 3H), whereas this preference was lost in 6-mo-old mutants by the fifth day (Fig. 3I), suggesting a marked decay in memory retrieval in older mutant animals.

Taken together, the findings from the Morris water maze test support an essential role for Wnt5a in the acquisition of spatial learning and memory storage in adult animals. Notably, the cognitive dysfunction in 3-mo-old CaMKII-Wnt5a^fl/fl mice were consistent with the LTP defects observed at this age but appeared before the onset of anatomical impairments. The more pronounced behavioral defects in 6-mo-old CaMKII-Wnt5a^fl/fl mice suggest a progressive decline in cognitive functions with the manifestation of dendritic abnormalities.

Wnt5a Loss Disrupts Calcium and Cytoskeletal Signaling Pathways and CREB-Mediated Transcription of Glutamate Receptors.

Wnts are known to exert their effects by signaling through three effector pathways: the canonical β-catenin–dependent pathway, a Ca²⁺-dependent pathway, and the planar cell polarity pathway (47). We found comparable levels of nuclear β-catenin and Axin2, c-myc, and NeuroD1, all transcriptional targets of canonical β-catenin signaling (48), between CaMKII-Wnt5a^fl/fl and control Wnt5a^fl/fl hippocampus at 3 mo (Fig. S5 A–C), indicating that canonical Wnt signaling is unaffected by Wnt5a depletion in the mature hippocampus.

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Fig. S5.

Wnt5a deletion perturbs calcium and planar cell polarity signaling, but not canonical Wnt signaling in the mature hippocampus. (A and B) Similar levels of nuclear (active) β-catenin in hippocampal nuclear fractions from CaMKII-Wnt5a^fl/fl (KO) mice and control littermates (WT) at 3 mo. The β-catenin immunoblot was reprobed for p84, a nuclear protein, for normalization. Results are means ± SEM from five mice per genotype. (C) Axin2, c-myc, and NeuroD1 transcript levels are unaffected by Wnt5a deletion in 3-mo-old KO hippocampus. Results are means ± SEM from six mice per genotype. (D) Representative immunoblots of synapse-associated signaling and structural proteins. Immunoblots were stripped and reprobed for tubulin for normalization. (E) Densitometric quantification of immunoblots of synaptic proteins in hippocampal homogenates from 3-mo-old KO and WT mice. Densitometric signal intensities for each protein were normalized to tubulin, and values expressed relative to control homogenates.

We next assessed the Wnt–calcium pathway, where Wnt ligands promote an increase of cytoplasmic Ca²⁺ (49, 50). Strikingly, Wnt5a treatment acutely elicited a calcium response in 92% of cultured rat hippocampal neurons transfected with GCaMP3, whereas only 43% of neurons responded to control treatment. Furthermore, the number of calcium transients was fivefold higher in Wnt5a-treated neurons (Fig. 4 A–C and Movies S1 and S2). We then performed biochemical analyses to assess activation of CaMKII, a critical regulator of hippocampal connectivity and functions (51, 52), in vivo, in young adult CaMKII-Wnt5a^fl/fl mice at 3 mo before the appearance of structural anomalies. Using a phospho-specific antibody that detects activated CaMKII (threonine 286 phosphorylation on CaMKIIα and T287 on CaMKIIβ) (53), we found a pronounced attenuation of phosphorylated CaMKIIα (59% decrease) and CaMKIIβ (57% decrease) in CaMKII-Wnt5a^fl/fl mice (Fig. 4 D and E). CaMKII-mediated phosphorylation of the GluA1 subunit of AMPA-type glutamate receptors at a critical serine 831 site (54, 55) has been functionally linked to synaptic plasticity and retention of spatial memory in mice (56, 57). We found a marked decrease in phospho-S831-GluA1 in postsynaptic density fractions from the mutant hippocampus (Fig. 4 D and E). These results suggest decreases in phosphorylation of CaMKII and GluA1 as the molecular underpinnings for the impairments in synaptic plasticity and spatial memory in CaMKII-Wnt5a^fl/fl mice.

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Fig. 4.

Wnt5a loss disrupts calcium and cytoskeletal signaling and decreases CREB-mediated GluN1 synthesis. (A) Wnt5a elicits a robust increase in calcium transients in hippocampal neurons. Rat hippocampal neuron cultures were transfected with GCaMP3 and treated with control L- or Wnt5a-conditioned media for 30 min. (Scale bar, 20 μm.) (B) Representative traces of calcium transients. (C) Wnt5a elicits a fivefold increase in calcium transients compared with control media. Results are mean ± SEM from n = 3 independent experiments (total of n = 31 L media-treated cells and n = 26 Wnt5a-treated neurons); ***P < 0.001, two-tailed t test. (D and E) Phosphorylated CaMKIIα/β and phosphorylation of GluA1^Ser831 are significantly reduced in CaMKII-Wnt5a^fl/fl (KO) hippocampus compared with Wnt5a^fl/fl litter-mates (WT) at 3 mo. Immunoblots were stripped and reprobed for total CaMKIIα, -β, and GluA1 for normalization. Results are mean ± SEM from n = 6 mice per genotype; *P < 0.05, ***P < 0.001, two-tailed t test. (F and G) CaMKIV and CREB phosphorylation are attenuated in hippocampal nuclear fractions from 3-mo-old KO mice. Blots were reprobed for total CaMKIV and CREB. Results are mean ± SEM from n = 6 and n = 5 mice per genotype; ***P < 0.001, two-tailed t test. (H and I) GluN1, the obligatory NMDA receptor subunit, is significantly decreased in 3-mo KO hippocampus, but levels of other NMDA receptor subunits, GluN2a/2b, and AMPA-type glutamate receptor subunits, GluA1 and GluA2, are unaltered. Immunoblots were stripped and reprobed for tubulin. Results are mean ± SEM from n = 6 mice per genotype; ***P < 0.001, two-tailed t test. (J) qPCR analysis shows decrease in GluN1 but not GluN2a/b transcripts in the KO hippocampus. Results are mean ± SEM from n = 6 mice per genotype; **P < 0.01, two-tailed t test. (K) A 1-kb region upstream of the mouse GluN1 promoter harbors three putative CRE sites. Hippocampal neurons were cotransfected with luciferase reporter constructs, Cypridina luciferase driven by the GluN1 promoter region, and Gaussia luciferase driven by an SV40 promoter as a control. Neurons were treated with control or Wnt5a media for 6 h and then harvested to measure luciferase activity. (L) Wnt5a treatment significantly increases GluN1 promoter-driven luciferase activity. Deletion of all three CRE sites or just the two proximal binding sites (CRE1 and CRE2) alone abolishes the Wnt5a-mediated response. Results are mean ± SEM for n = 6 independent experiments; *P < 0.05, **P < 0.01, ***P < 0.001, two-tailed t test. (M and N) Rac1-GTP levels are reduced in 3-mo KO mice. Hippocampal homogenates were subjected to GST-PAK-PBD-agarose pull-downs for active Rac-GTP and immunoblotted for Rac1. Rac1-GTP signal intensities were normalized to total Rac1. Results are mean ± SEM from n = 6 mice per genotype; **P < 0.01, two-tailed t test. (O and P) Phospho-JNK levels are significantly decreased in hippocampal homogenates from 3-mo KO mice. Results are means ± SEM from n = 6 mice per genotype; *P < 0.05, **P < 0.01, two-tailed t test.

Calcium signaling within synapses could couple to transcriptional responses via shuttling of a Ca^2+/CaM/CaMKIIγ complex to the nucleus to promote phosphorylation of CaMKIV, which then phosphorylates and activates the transcription factor CREB (58). Phosphorylation of CaMKIV and CREB were significantly reduced in nuclear fractions from 3-mo-old CaMKII-Wnt5a^fl/fl hippocampal tissues (Fig. 4 F and G). Because Wnt5a deletion altered nuclear CREB phosphorylation, we assessed levels of several synaptic proteins (Fig. S5 D and E) and found that only GluN1, the obligatory NMDA receptor subunit, was decreased (Fig. 4 H–J), raising the possibility that GluN1 transcription is CREB-dependent. We did not observe any changes in levels of other NMDA receptor subunits, GluN2a/2b, that are coexpressed and coassembled in the endoplasmic reticulum (ER) with GluN1, in the mature hippocampus (Fig. 4 H–J). We identified three putative CRE sites (CRE1 at −212 bp, CRE2 at −238 bp, and CRE3 at −770 bp) in a 1-kb region upstream of the transcription start site in the mouse GluN1 promoter (Fig. 4K). In a dual luciferase assay, Wnt5a stimulation of hippocampal neurons for 6 h significantly increased luciferase activity compared with control treatment (Fig. 4 K and L). Mutation of just the two proximal CRE elements (−212 to −216 bp and −238 to −242 bp) abolished Wnt5a-induced luciferase activity (Fig. 4L). These results reveal an unexpected role for Wnt5a in enhancing GluN1 transcription through a noncanonical pathway that involves calcium-CaMKII-CREB activation.

We finally examined the planar cell polarity pathway where noncanonical Wnts induce cytoskeletal dynamics by activating small GTPases, such as Rac1 and JNK signaling (59). Rac1 is a critical regulator of the actin cytoskeleton in dendrites and spines (60, 61). Active Rac1-GTP and phospho-JNK levels were significantly reduced in hippocampal homogenates prepared from 3-mo-old CaMKII-Wnt5a^fl/fl mice (Fig. 4 M–P). Rac1 activity can also be influenced by CaMKII activity via CaMKII-mediated phosphorylation of the Rac1-specific GEFs, Tiam1 and Kalirin-7 (62, 63). Taken together, these results suggest that Wnt5a signals via CaMKII and Rac1-mediated signaling, as well as CREB-mediated GluN1 synthesis to maintain synaptic plasticity and structure in the adult hippocampus.

Reagents and Antibodies.

The antibodies used in this study were previously validated for the following applications: Wnt5a (R&D Systems; AF645, Western blotting), VGLUT1 (Millipore; AB5905, Western blotting), PSD95 (NeuroMab; Western blotting), GluA1 [Johns Hopkins Medical Institutions (JHMI) monoclonal antibody core; Western blotting], GluA2 (JHMI monoclonal antibody core; Western blotting), GluN1 (Millipore; MAB363, Western blotting), CaMKII (Cell Signaling; 4436, Western blotting), Tubulin (Sigma Aldrich; T-6199, Western blotting), Rac1 (Millipore; 05-389, Western blotting), P-CaMKII (Thr286) (Cell Signaling; 3316 and 12716, Western blotting), P-CaMKIV (Thr196) (Santa Cruz; sc-28443-R, Western blotting), CaMKIV (Abcam; ab3557, Western blotting), P-CREB (Ser133) (Cell Signaling; 9198, Western blotting), CREB (Cell Signaling; 9104, Western blotting), GluN2a (JHMI monoclonal antibody core; Western blotting), GluN2b (JHMI monoclonal antibody core; Western blotting), Akt (Cell Signaling; 9272, Western blotting), Erk (Cell Signaling; 9102, Western blotting), Nuclear matrix protein p84 (GeneTex; GTX70220, Western blotting), Active β-catenin (Millipore; 05-665, Western blotting), p-JNK (Cell Signaling; 9251, Western blotting), JNK (Cell Signaling; 9252, Western blotting), p-GluA1-Ser831 (Millipore; AB5847, Western blotting), NeuN (Millipore; MAB377, immunohistochemistry), Neurofilament (Millipore; AB5539, immunohistochemistry), MAP2 (Sigma Aldrich; M9942, immunohistochemistry) Synaptophysin (Sigma Aldrich; S5768, Western blotting) and Synapsin 1 (BD Transduction laboratories; 611392, Western blotting). Golgi-Cox staining was done using the FD Rapid GolgiStain kit (FD Neurotechnologies; PK401). Rac1/Cdc42 GTPase activity was measured by a kit (EMD Millipore; 17-441). GluN1 promoter activities were followed with BioLux Cypridina and Gaussia luciferase assay kits (New England Biolabs, E3309S and E3300S). Wnt5a conditioned media and L-cell conditioned media were harvested from Wnt5a producing cells (ATCC CRL-2814) and L cells (ATCC CRL-2648). Both cell lines were first cultured in DMEM supplemented with 10% (vol/vol) FBS for 2 d, and then the medium was changed to Neurobasal medium supplemented with B27 for 24 h.

In Situ Hybridization.

In situ hybridization was performed using a digoxigenin-labeled probe spanning a 572-bp region within Wnt5a exon 2. Mouse brains of various ages were fresh frozen in OCT (Tissue-Tek) and serially sectioned (20 μm) using a cryostat. Sections were postfixed in 4% paraformaldehyde (PFA), washed in PBS, and acetylated with 0.25% acetic anhydride in 0.1 M triethanolamine with 0.9% NaCl. After hybridization with the labeled RNA probe (2 μg/mL) at 68 °C overnight, sections were washed with 0.2× SSC buffer at 65 °C, blocked with TBS containing 1% normal goat serum, and then incubated with alkaline phosphatase-labeled anti-DIG antibody (1:5,000; Roche) overnight at 4 °C. The alkaline phosphatase reaction was visualized with NBT/BCIP, rinsed in PBS, fixed in 4% PFA and mounted in AquaMount (EMD Chemicals).

Real Time-PCR Analyses.

Total RNA was prepared from dissected hippocampal tissues using TRIzol-chloroform extraction (ThermoFisher; 10296-010). RNA was then reverse-transcribed using a RETROscript Reverse Transcription kit (ThermoFisher; AM1710). Real-time qPCR was performed using a Maxima SYBR Green/Rox Q-PCR Master Mix (ThermoFisher; K0221), in 7300 Real time PCR System (Applied Biosystems). The following primers were used for the analyses; Wnt5a-F:5′-CTCGGGTGGCGACTTCCTCTCCG-3′ and Wnt5a-R:5′-CTATAACAACCTGGGCGAAGGAG-3′; Wnt2-F:5′-GTAGATGCCAAG GAGAGGAAAG-3′ and Wnt2-R:5′-CCACTCACACCATGACACTT-3′; Wnt3-F:5′-TGGACCACATGCACCTAAAG-3′ and Wnt3-R:5′-CGTACTTGTCCTTGAGGAAG TC-3′; Wnt3a-F:5′-GCAGCTGTGAAGTGAAGAC-3′ and Wnt3a-R:5′-GGTGTTTCT CTACCACCATCTC-3′; Wnt5b-F:5′-GAGAGCGTGAGAAGAACTTTG-3′ and Wnt5b-R: 5′-GCGACATCAGCCATCTTATAC-3′; Wnt7a-F: 5′-GCCTTCACCTATGCGAT TATC-3′ and Wnt7a-R: 5′- GGTACTGGCCTTGCTTCTC-3′; Wnt7b-F: 5′-GCATGA AGCTGGAATGTAAGTG-3′ and Wnt7b-R: 5′-TGCGTTGTACTTCTCCTTGAG-3′; Wnt8a-F: 5′-TGGGAACGGTGGAATTGTC-3′ and Wnt8a-R: 5′-GCGGATGGCAT GAATGAAG-3′; Wnt11-F: 5′-CCTGGAAACGAAGTGTAAATGC-3′ and Wnt11-R: 5′-TGACAGGTAGCGGGTCTTG-3′; GluN1-F: 5′-CCAGATGTCCACCAGACTAA AG-3′ and GluN1-R: 5′-CATTGACTGTGAACTCCTCTTTG-3′; GluN2a-F: 5′-CTGTGTGGCCAAGGTATAAG-3′ and GluN2a-R: 5′-TCAGTCAGTGGGTCTAT GTC-3′; GluN2b-F: 5′-ATGAGGAACCAGGCTACATC-3′ and GluN2b-R: 5′-GGT CACCAGGTAAAGGTCATAG-3′; Axin2-F: 5′-GAGAGTGAGCGGCAG AGC-3′ and Axin2-R: 5′-CGGCTGACTCGTTCTCCTG-3′; NeuroD1-F:5′-GCTACTCCAAGACC CAGAAAC-3′ and NeuroD1-R:5′-TGTACGAAGGAGACCAGATCA-3′; c-myc-F: 5′-CTCCGTACAGCCCTATTTCATC-3′ and c-myc-R: 5′-TGGGAAGCAGCTCGAATT TC-3′; GAPDH-F: 5′-CCTGCACCACCAACTGCTTA-3′ and GAPDH-R: 5′-CC ACGATGCCAAAGTTGTCA-3′. Each sample was analyzed in triplicate reactions. Data were analyzed using the ΔΔC_t method, normalizing each sample to the internal control, and relative messenger RNA was determined as the percentage of the maximum value observed in the experiment.

Immunohistochemical Analyses.

Mouse brains were fixed in 4% PFA at 4 °C overnight, cryoprotected in 30% sucrose in PBS, frozen in OCT (Tissue-Tek) with dry ice, and serially sectioned (40 μm). For immunohistochemistry with diaminobenzidine (DAB), sections were first incubated with 10% methanol + 3% H₂O₂ for 20 min to quench peroxidase activity and then washed in TBS, permeabilized in TBS containing 0.4% Triton X-100, and blocked using 10% goat serum + 3% BSA for 2 h. Sections were incubated in the following primary antibodies overnight: chicken antineurofilament (EMD Millipore; AB5539, 1:500) or mouse anti-MAP2 (Sigma Aldrich; M9942, 1:1,500). Following TBS washes, sections were incubated with either chicken or mouse anti-HRP secondary antibodies (GE Healthcare; 1:500) for 1 h. Sections were then washed in TBS, incubated with DAB for 2–10 min at room temperature, followed by washes in TBS and then mounted in VectaShield (Vector Laboratories).

For immunofluorescence, sections were washed in TBS, permeabilized in TBS containing 0.4% Triton X-100, and blocked using 10% goat serum + 3% BSA for 2 h. Sections were then incubated in primary antibodies overnight. Following TBS washes, sections were incubated with Alexa-Fluor conjugated secondary antibodies (ThermoFisher; 1:500) for 1 h. Sections were washed in TBS and then mounted in VectaShield (Vector Laboratories).

Neuronal Cell Counts.

Six- and 12-mo mouse brains for neuronal counts were prepared as described in Ramanan et al. (88). Briefly, mouse brains were fixed in PBS containing 4% PFA, and then cryoprotected overnight in 30% sucrose-PBS. Coronal brain sections (20 μm) were stained with a solution containing 0.5% Cresyl violet (Nissl), and cells in CA1, CA3, and dentate granule layers were counted at 40× magnification. For each mouse, 9–12 defined regions were counted and analyzed from three coronal sections using ImageJ.

Viral Injections.

Lentiviruses expressing GFP or GFP-T2A-CRE were generated by subcloning into lentiviral backbones using XhoI and NotI restriction sites. Lentivirus was produced in HEK293T cells using the FUW/Δ8.9/VSVG system. Two collections of media were done 24 and 48 h after transfection, and virus particles pelleted by ultracentrifugation. Virus particles were then resuspended in Neurobasal media and stored at –80 °C until use.

AAV-EF1a-DIO-mCherry and AAV-CMV-GFP were obtained from University of North Carolina Vector Core (2.4 × 10¹³ viral particles per milliliter, stock by Deisseroth laboratory) and University of Pennsylvania Vector Core (5.2 × 10¹³ viral particles per milliliter), respectively. AAV-CMV-DIO-Wnt5a was generated by Vigene Biosciences (2.8 × 10¹³ viral particles per milliliter).

For viral injections, mice were deeply anesthetized with avertin (tribromoethanol, 0.25 mg/g body weight; 2–methyl–2–butanol, 0.16 μL/g body weight) and mannitol (to prevent edema, 10 mg/g body weight). Lentiviral or AAV vectors were stereotaxically delivered to the hippocampus using coordinates that were previously described (89). After injections, mice were allowed to recover and killed for biochemical analyses 2 wk after infections for biochemical analyses or 3 mo later for morphological analyses.

Dendrite Reconstructions and Analyses.

Thy1-GFP mouse brains were harvested and fresh frozen in OCT, and serial coronal sections (200 μm) were prepared using a cryostat. Brain sections were mounted on slides and preserved with PermaFluor mounting medium (Thermo Scientific). GFP-filled CA1 pyramidal neuronal morphologies were imaged using 20× objective to visualize entire dendritic fields (512 × 512 pixels; 1-μm optical scanning) followed with a 3 × 4 tile scan using a Zeiss LSM 710 confocal microscope (Carl Zeiss). Z-stacks were stitched together using Zen imaging software. All of the analyzed neurons were sufficiently bright to allow complete tracing of dendrite arbors, and had intact dendritic arbors. Neuronal somas and dendritic structure were manually traced with Imaris 7.7 software (Bitplane) to create 3D reconstructions of dendritic structures. Dendritic arbors that could not be confidently reconstructed were not used in the analyses. Total dendrite lengths were quantified using Imaris. Dendritic complexities were analyzed by 3D Sholl analysis, and calculating the number of dendrite intersections with concentric spheres that radiated from the soma in 10-μm-radius increments. Five neurons were traced per animal, and the average used as the value for each mouse during statistical analyses.

Golgi staining was performed with FD Rapid Golgistain kit on mouse brains that were fresh-frozen in OCT and serially sectioned (200 μm) using a cryostat. Dendritic arbors were analyzed as described previously (90). Briefly, Golgi-impregnated pyramidal neurons in the CA1 region were traced using Neurolucida software (MicroBrightField) under a Nikon Eclipse E800 microscope, equipped with a motorized stage. Analyses were done by an investigator blinded to the genotype. All of the analyzed neurons were well stained, isolated and had intact dendritic arbors. Dendritic length of each traced neuron was calculated using NeuroExplorer software (MicroBrightField). Five neurons were traced per animal, and the average used as the value for each mouse during statistical analyses.

To measure spine density, we took images of distal apical dendrites (>100 μm away from the soma) of hippocampal CA1 pramidal neurons at 63× magnification using a Zeiss Axiovision microscope with a AxioCam HRC digital camera. The position of each dendritic spine along these dendrite segments was assessed manually and counted using ImageJ software. Spine density for each animal was obtained from at least 15 dendrites per mouse with a total length of 2,000~3,000 μm traced for statistical analysis.

Rac1 GTPase Activity.

The Rac1/Cdc42 Activation Assay (EMD Millipore; 17–441) was used to detect Rac1 activity in hippocampal homogenates. Briefly, hippocampal lysates were precleared with glutathione-agarose beads (100 μL). Supernatants were incubated with a fusion protein (GST-PAK) coupled to agarose beads. GST-PAK specifically binds to Rac1 or Cdc42 in its GTP-bound form. Rac1-GTP in the pull-downs was detected by immunoblotting using a Rac1-specific antibody.

Slice Preparations and Electrophysiology.

Three- or 6-mo-old male mice were anesthetized with the inhalation anesthetic isoflurane before decapitation. Brains were rapidly dissected out and placed in ice-cold, oxygenated (95% O₂ and 5% CO₂) low-Ca²⁺/high-Mg²⁺ dissection buffer containing 2.6 mM KCl, 1.25 mM NaH₂PO₄, 26 mM NaHCO₃, 211 mM sucrose, 11 mM glucose, 0.5 mM CaCl₂, and 7 mM MgCl₂. The 350-μm transverse hippocampal slices were prepared using a vibratome (Leica; VT1000s) in dissection buffer followed by recovery in a static submersion chamber filled with oxygenated artificial cerebrospinal fluid (ACSF) containing 125 mM NaCl, 5 mM KCl, 1.25 mM NaH₂PO₄, 2 mM CaCl₂, 1 mM MgCl₂, 25 mM NaHCO₃, and 11 mM glucose at 30 °C for at least 1 h before LTP or 2 h before LTD recording.

Slices were transferred to a recording chamber where they were perfused continuously with oxygenated ACSF at a flow rate of ~3 mL/min at 30 °C. Hippocampal CA1 fEPSP was evoked at 0.033 Hz with a 125-μm platinum/iridium concentric bipolar electrode (FHC) placed in the middle of the stratum radiatum of CA1. Synaptic responses were recorded with ACSF-filled microelectrodes (1–2 MΩ), positioned ~200 μm away (orthodromic) from the stimulating electrode, and were quantified as the initial slope of fEPSP. Input–output relationships were obtained for each slice with various stimulus intensity and responses were set to ~45% maximum for LTP experiments and ~55% maximum for LTD experiments. Input–output curves were plotted as the fEPSP slope against the presynaptic fiber volley amplitude across all stimulation intensities and the slope for the linear fit of the fiber volley–fEPSP slope relationship were obtained from each slice. PPF was recorded with an interstimulus interval of 25−250 ms. PPF data were presented as a ratio of the second response slope relative to the first. LTP was induced by θ-burst stimulation, consists of four trains of 10 bursts at 5 Hz, with each burst consisting of four stimuli given at 100 Hz and 10-s intertrain interval. LTD was induced by long-frequency stimulation consists of 900 single pulses at 1 Hz.

All hippocampal slice electrophysiological recordings were performed by an experimenter blind to the animal genotype. Statistical significance was determined with a two-way ANOVA for PPF and a two-tailed, unpaired t test for input–output relationship, LTP, and LTD. Representative traces are averages of 4 (PPF and input–output relationship) or 10 (LTP and LTD) traces, and stimulus artifacts have been removed for clarity.

Calcium Imaging.

Hippocampal neurons from E18 rat pups were plated onto glass coverslips coated with poly-l-lysine (1 μg/mL; Sigma-Aldrich) and grown in Neurobasal growth medium supplemented with 2% B27, 2 mM GlutaMax, 50 U/mL penicillin, 50 μg/mL streptomycin, and 5% FBS. Neurons were switched to serum-free Neurobasal medium 24 h postseeding and fed twice a week. Neurons were cotransfected with GCaMP3 and TdTomato at 14–15 d in vitro using lipofectamine 2000 (Invitrogen). Briefly, coverslips containing neurons were assembled onto a closed perfusion chamber and continuously perfused with ACSF buffer with 2 mM Mg²⁺. After 10 min of baseline recording (F₀), neurons were perfused with Wnt5a- or L-conditioned media diluted (1:500) in ACSF buffer with 1 mM Mg²⁺ for 30 min. All imaging experiments were performed at 37 °C using a Zeiss spinning disk confocal (Carl Zeiss). The GCaMP3 was imaged at 488-nm excitation and collected through a 505- to 550-nm filter, whereas the TdTomato signal was imaged at 561-nm excitation and 575- to 615-nm emission. Neurons were imaged using a 63× oil objective (N.A. = 1.40) at a rate of six images per minute. Images were analyzed using ImageJ software (NIH) by calculating the normalized change in average GCaMP3 over TdTomato fluorescence intensities from neuronal soma. The fluorescence intensity change is expressed as ΔF/F₀ and the amplitude of fluorescence change (ΔF_max/F₀) represents the extent of GCaMP3 intensity change.

Luciferase Assays.

To monitor GluN1 promoter activity, the GluN1–Cypridina reporter construct was generated by subcloning a 1-kb region upstream of the transcription start site of mouse GluN1 into pCLuc Mini-TK 2 Vector (New England Biolabs). Neurons were cotranfected with GluN1–Cypridina reporter and a control pSV40-GLuc vector that encodes for Gaussia luciferase under the control of the constitutive SV40 promoter. Two days following transfection, neurons were stimulated with Wnt5a or L media diluted 1:500 in culture media. Because the luciferase proteins are secreted, supernatants were harvested and mixed with either Cypridina or Gaussia luciferase substrates and reporter gene activity was monitored with BioLux Cypridina and Gaussia luciferase assay kits (New England Biolabs) using a luminometer.

We thank Hey-Kyoung Lee, Gareth Thomas, Naoya Yamashita, and Xin Chen for insightful comments on the manuscript; Vince Hilser for providing luciferase constructs; Eric Wang and Huaqiang Fang for assistance with immunohistochemistry and calcium imaging, respectively; and Michelle Pucak at the Multiphoton Imaging Core (The Johns Hopkins University School of Medicine) and Erin Pryce at the Integrated Imaging Center (The Johns Hopkins University) for help with imaging and Imaris analyses. This work was supported by NIH Grants NS073751 (to R.K.), NS073930 (to B.X.), DC007395 (to H.Z.), GM076430 and EY024452 (to S.H.), and NS036715 (to R.L.H.). C.-M.C. was supported, in part, by NIH Training Grant T32GM007231 to the Biology Department.