Lnx1^−/− (Liu et al., 2018) knockout and EphB2^F620D (Holmberg et al., 2006) knockin mice and genotyping methods have been described previously. Lnx1^−/− mice were crossed with the Thy1-GFP M (Feng et al., 2000) transgenic mouse line, which were generously provided by Mark Henkemeyer (University of Texas Southwestern Medical Center, Dallas, TX, USA). Mice were anesthetized with isoflurane and perfused with 0.1 M phosphate-buffered saline (PBS) followed by 4% paraformaldehyde in phosphate buffer. The brains were then removed, postfixed, and sectioned at 30 μm using a vibratome. All experiments that involved mice were carried out in accordance with the US National Institutes of Health Guide for the Care and Use of Animals under an Institutional Animal Care and Use Committee approved protocol and Association for Assessment and Accreditation of Laboratory Animal Care approved Facility at the Shanghai Jiao Tong University School of Medicine. Mice were raised in animal facilities with a constant temperature (22°C) and on a 12-h light-dark cycle. Food and water were unlimited to access.

For immunofluorescence, vibratome sections were blocked with permeable buffer (0.3% Triton X-100 in PBS) that contained 10% donkey serum for 30 min at room temperature, incubated with primary antibodies in a permeable buffer that contained 2% donkey serum overnight at 4°C. The slices were then washed three times with PBS-T (0.1% Tween-20 in PBS) for 10 min every time and incubated with Alexa Fluor secondary antibodies (1:200, Molecular Probes) in the PBS buffer for 2 h at room temperature. The slices were washed in PBS-T three times and mounted on glass slides using Aqua-Poly/Mount (Polysciences) and photographed using a confocal microscope (Leica SP8). For primary antibodies, we used goat anti-EphB2 (1:500, R&D, P54763), rabbit anti-synapsin1 (1:1,000, gift from Ilya Bezprozvanny, University of Texas Southwestern Medical Center, Dallas, TX, USA), and rabbit anti-proteasome S20 (1:1,000, Abcam, ab3325).

To quantify the shape of the spine, a procedure was adapted from our previous study (Xu et al., 2011). To classify the shape of neuronal spines in slices, we used the NeuronStudio software package and an algorithm with the following cutoff values: aspect ratio (AR)_thin (crit) = 2.5, head to neck ratio (HNR) (crit) = 1.3, head diameter (HD) critical value (crit) = 0.4 μm. The protrusions with length 0.2–3.0 μm and Max width 3 μm were counted. Spine density was calculated by dividing the total spine number by the dendritic branch length. Primary dendrite was defined as the dendrite from the soma. The localizations of the terminals and spines were carefully identified in the 3D rendering views. The synapsin1⁺ spines were defined as the overlapped connections of spines and synapsin1⁺ terminals with any identical red/green pixels, otherwise synapsin1⁻ (separated from terminals). The neurons of the CA3 corner area were chosen and spines in the stratum oriens layer of the CA3 area were counted. Control and experiment conditions were adjusted with the same parameters. Acquisition of the images and morphometric quantification were performed under “blinded” conditions.

Mice were perfused with 2.5% glutaraldehyde in phosphate buffer, pH 7.2 for 30 min, dissected brains were then postfixed in the same buffer overnight at 4°C. After PBS buffer rinse, samples were postfixed in 1% osmium tetroxide buffer (2 h) on ice in the dark. Following a double-distilled water rinse, tissue was stained with 3% aqueous uranyl acetate (0.22 μm filtered, 1 h dark), dehydrated in a graded series of ethanol, propylene oxide, and embedded in Epoxy 618 resin. Samples were polymerized at 60°C for 48 h. Thin sections, 60–90 nm, were cut with a diamond knife on the LKBV ultramicrotome and picked up with Formvar-coated copper slot grids. Grids were stained with lead citrate and observed with transmission microscopy (PHILIP CM-120). Images from the CA3 corner area were chosen and an average of 8 pictures per mouse (n = 4 animals per genotype) were captured. The PSDs or PSD length was counted or analyzed by Image J software under “blinded” conditions.

Primary cell culture of hippocampal neurons was performed as described (Xu and Henkemeyer, 2009). Briefly, hippocampal neurons were dissociated from postnatal day 0 (P0) pups. The triturated cells (1 × 10⁵ cells per well) were grown on either 6 well dishes or glass coverslips coated with 10 μM polylysine overnight in 24 well dishes. Then the culture was grown in a medium of Neurobasal A media (GIBCO) supplemented with B27 and 2 mM glutamine for indicated days.

For receptor trafficking assay, cultured neurons were incubated with special N-terminus EphB2 antibodies (1:100, R&D, P54763) for 10 min. After three washes with preheated PBS, Alexa 488-conjugated secondary antibody (1:100) in blocking buffer was then applied for 10 min to label the total membranous receptors. After a rapid rinse with culture medium, neurons were placed back in the incubator with the original culture medium for 30 min to allow the trafficking of receptors. Then EphB2 antibodies (1:100) followed by Alexa 555-conjugated secondary antibody were applied for 10 min to label the trafficking receptors. After three washes with PBS, the neurons were fixed in 4% paraformaldehyde for 10 min and mounted.

For receptor internalization assay, cultured neurons were incubated with special N-terminus EphB2 antibodies (1:100, R&D, P54763) for 10 min. After a rapid rinse with a culture medium, neurons were placed back in the incubator for 30 min to allow the internalization of receptors. Alexa 488-conjugated secondary antibody (1:100) in blocking buffer was applied for 10 min to label the existing surface EphB2 receptors. After three washes with preheated PBS, the neurons were fixed in 4% paraformaldehyde for 10 min and permeabilized with a blocking buffer that contained 0.1% Triton-X 100 for 30 min. Alexa 555-conjugated secondary antibodies were then applied for 10 min to label the internalized population of receptors. Finally, the neurons were further incubated with proteasome S20 antibody (1:1,000, Abcam, ab3325) in a permeable buffer that contained 2% donkey serum overnight at 4°C and corresponding Alexa 647-conjugated secondary antibody (1:1,000) on the next day. After three washes with PBS, the neurons were mounted.

Primary hippocampal neurons from wild-type (WT) or Lnx1^−/– pups were cultured for 3 days, then GFP or FLAG-Lnx1 virus (OBIO Company, Shanghai, China) was added into the neurons for another 7-day culture. Then the primary hippocampal neurons were prepared for cell surface protein isolation and western blotting. Cell surface protein isolation of the cultured primary hippocampal neuron samples was separated using a Pierce Cell Surface Protein Isolation Kit (Thermo) as the manufacturer's protocol. Total and membrane proteins were separated by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE), transferred to nitrocellulose membranes, and then immunoblotted with indicated antibodies. Analysis of the data was performed using NIH ImageJ software, the mean density of each band was normalized to β-actin signal in the same sample and averaged. For primary antibodies, we used mouse anti-β-actin (1:3,000, Thermo, MA5-15739), mouse anti-Flag (1:1.000, Sigma, F1804), and Goat anti-EphB2 (1:1,000, R&D, P54763).

Brain coronal slices were prepared from 3-week-old naive Lnx1^+/+, Lnx1^−/–, EphB2^F620D, and Lnx1^−/–; EphB2^F620D mice. Brains were dissected quickly and chilled in ice-cold artificial cerebrospinal fluid (ACSF) that contains (in mM): 125 NaCl, 2.5 KCl, 2 CaCl₂, 1 MgCl₂, 25 NaHCO₃, 1.25 NaH₂PO₄, and 12.5 Glucose. Coronal brain slices (300 μm thick) were prepared with a vibratome and recovered in ACSF bubbling with 95% O₂ and 5% CO₂ at 31°C for 1 h and then maintained at room temperature (22–25°C). The electrode internal solution was composed of 115 mM CsMeSO₃, 10 mM HEPES, 2.5 mM MgCl₂·6H₂O, 20 mM CsCl₂, 0.6 mM ethylene glycol tetraacetic acid (EGTA), 10 mM Na₂phosphocreatine, 0.4 mM Na-guanosine 5′-triphosphate (GTP), and 4 mM Mg-adenosine triphosphate (ATP). For mEPSC recording, tetrodotoxin (1 μM) and picrotoxin (100 μM) were included in the external solution. Cells were held at −70 mV. Miniature responses were acquired with a Multiclamp 200B at 10 kHz. Prior to mEPSC detection and analysis, current traces were low-pass filtered at 5 kHz. Data were analyzed in pClamp 10.6 (Molecular Devices) and recordings were made from an average of 3 cells per slice and 2–3 slices per mouse.

The results are presented as mean ± SEM. Statistical differences were determined by Student's t-test for two-group comparisons or ANOVA followed by Turkey test for multiple comparisons among more than two groups.

We have identified that Lnx1 was expressed especially in the postsynaptic fraction of hippocampal CA3 neurons (Liu et al., 2018), the postsynaptic localization prompted us to detect whether neuronal morphogenesis is altered in Lnx1^−/− mice. We compared the dendrites and spines between postnatal week 3 (PW3) Lnx1^−/− and WT mice with a Thy1-GFP-M transgenic reporter (Feng et al., 2000) and found a higher spine density of CA3 but not CA1 pyramidal neurons in Lnx1^−/− mice than that of WT littermate mice (Figures 1A,B), though no difference was observed in primary dendrites (data not show). Analysis of spine morphology showed that the spine head width and spine length in CA3 pyramidal neurons of Lnx1^−/− mice were significantly decreased (Figures 1A,C). These results indicated that the deficiency of lnx1 in CA3 neurons may lead to changes in the morphology and function of dendritic spines. To examine whether the changes in spine density and morphology resulted in changes in synapse number or postsynaptic structure, we further performed transmission EM and analyzed the number of synapses in the CA3 area and their postsynaptic morphology in PW3 mice. We observed a significant increase in the density of synapses with a decreased postsynaptic profile area indicated by its PSD length in Lnx1^−/− mice as compared to WT mice (Figures 1D–G). These results demonstrate a critical role of Lnx1 in the process of spinogenesis in adolescent mice.

The abnormal spine morphogenesis in Lnx1^−/− mice promotes us to ask if the structure of synapses is anatomically altered. Brain slices from PW3 WT and Lnx1^−/− mice with Thy1-GFP-M transgenic reporter were immunostained with anti-synapsin1, a protein marker for presynaptic terminals. In WT mice, most spines of CA3 neurons were contacted with presynaptic terminals (synapsin1⁺ spine), while this ratio was decreased in Lnx1^−/− mice (Figures 2A–C). We further classified the synapsin1⁺ spines of CA3 neurons into three types based on their morphology: one presynaptic terminal on one spine (Type I, typical 1:1 spine), two or more presynaptic terminals on one spine (Type II, multi-innervated spines), and one presynaptic terminal on two or more spines (Type III, multi-spine boutons) (Figures 2A,B). We observed a differential distribution of the three type synapses, Type I the highest and Type III the lowest, in WT mice. However, Lnx1^−/− mice showed an increased rate of Type III synapse and a decreased rate of Type II synapse (Figure 2D). To validate the phenotype observed by immunofluorescence staining, we performed transmission EM to analyze these synapse types (Figure 2E). We observed an increased rate of Type III synapse and a decreased rate of Type II synapse, while the percent of Type I synapse remains unchanged in Lnx1^−/− mice when compared with littermate controls (Figure 2G). We further classified the Type I synapse into two subtypes, continuous PSDs (Type I-A) and segmented PSDs (Type I-B) (Figure 2F), and observed a significantly increased ratio of Type I-B to Type I-A synapse in Lnx1^−/− mice (Figure 2H). Combined with our previous results that postsynaptic efficacy was greatly reduced in the CA3 neurons of Lnx1^−/− mice (Liu et al., 2021), these results thus demonstrate a critical role for Lnx1 to establish an intact postsynaptic structure that attributes to the formation of functional synapses in adolescence mice.

In the previous study, we screened molecules that were involved in synaptogenesis mediated by Lnx1 and found that the EphB receptors, a family of tyrosine receptor kinases that regulate excitatory synaptogenesis early during development (Sheffler-Collins and Dalva, 2012), were reduced in cell membrane component extracted from Lnx1^−/− hippocampus. Lnx1 stabilized EphB receptors through specific binding sites to prevent their degradation in the proteasome (Liu et al., 2018). Lnx1 knockout decreased the surface localization of EphB receptors, which could be the result of either impaired trafficking of EphBs receptors into the plasma membrane or enhanced internalization of EphBs from the plasma membrane. To further determine how Lnx1 regulated the surface expression of EphB receptors, we cultured Lnx1^−/− hippocampal neurons and investigated the surface trafficking of the EphB2 receptor. We firstly used a special antibody against the N-terminus of the EphB2 receptor and corresponding fluorescence secondary antibody (488, green) to label the already existing surface EphB2 receptors at DIV10 (day in vitro), after a 30 min trafficking, we used N-terminal EphB2 antibody again and another corresponding fluorescence secondary antibody (555, red) to label new surface EphB2 receptors that trafficked from cytoplasm to membrane in the 30-min period (Figure 3A). We observed the surface EphB2 receptors (488, green) were decreased in the Lnx1^−/− hippocampal neurons when compared with WT neurons, which was inconsistent with our previous results that loss of Lnx1 reduced surface EphB receptors (Figures 3B,C). However, the newly trafficked EphB2 receptors (555, red) showed no difference between WT and Lnx1^−/− neurons (Figures 3B,C), indicating that Lnx1 did not affect the trafficking of EphB receptors into the plasma membrane.

Then we investigated whether the internalization of EphB2 receptors from the plasma membrane was affected by Lnx1. Similarly, we used an antibody against the N-terminus of the EphB2 receptor to bind the surface EphB2 receptors and allow them to internalize from the plasma membrane. We added the corresponding fluorescence secondary antibody (488, green) to label the existing surface EphB2 receptors after 30-min incubation, then neurons were fixed and permeabilized for another corresponding fluorescence secondary antibody (555, red) to label the internalized EphB2 receptors from the plasma membrane (Figure 3D). We found that the ratio of internalized EphB2 was increased in Lnx1^−/− hippocampal neurons as compared to WT neurons (Figures 3E,F). Furthermore, we extracted the membrane fraction of cultured primary hippocampal neurons from WT or Lnx1^−/− pups and observed a decreased EphB2 level in both total and membrane fractions in Lnx1^−/− neurons, which could be restored to a comparable normal level after the overexpression of Lnx1 protein (Figure 3G). Together, these data suggest that Lnx1 is necessary and sufficient for the stability of EphB receptors by preventing their internalization from the cell surface.

We have found that Lnx1 stabilized EphB receptors by preventing their degradation in proteasome in our previous study (Liu et al., 2018). To test whether the internalized EphB2 receptors in Lnx1^−/− hippocampal neurons were subsequently submitted to proteasome for degradation, we performed internalization assays and immunofluorescence staining with anti-proteasome S20 to label the proteasomes. In WT neurons, we observed little co-localization between EphB2 receptors and proteasomes, while a portion of internalized EphB2 receptors in Lnx1^−/− neurons were co-localized with proteasome S20 protein (Figures 4A–C), suggesting that Lnx1 is necessary for arresting EphB2 receptor internalization for proteasomal degradation.

EphB2 forward signaling has been reported to be required for the spine morphogenesis and synapse formation in the CA3 pyramidal neurons in vivo (Henkemeyer et al., 2003). To determine whether constitutive catalytic activation of EphB2 is able to reverse the morphological defect caused by Lnx1 ablation, EphB2^F620D/F620D, a constitutively active form of EphB tyrosine kinase (Holmberg et al., 2006), was crossed with Lnx1^−/− mice. We first detected the ratio of internalized EphB2 receptors in WT, Lnx1^−/− and Lnx1^−/−; EphB2^F620D/F620D primary hippocampal neurons, and found that the increased ratio of internalized EphB2 in Lnx1^−/− was recovered in Lnx1 ^−/−; EphB2^F620D/F620D neurons (Figures 4A,B). Moreover, the ratio of co-localization between internalized EphB2 receptors and proteasomes was also restored to a comparable normal level in Lnx1^−/−; EphB2^F620D/F620D neurons (Figure 4C), which suggest that active EphB2 is more resistant to the degradation. We next examined the spine morphology of CA3 neurons and found that the EphB2^F620D/F620D mice per se remain unchanged when compared to the WT mice, while the shrunken spines in CA3 neurons were completely recovered in Lnx1^−/−; EphB2^F620D/F620D compound mutants (Figures 4D–F). We also examined the rearrangement of all types of synapses and found that the EphB2^F620D/F620D mice per se showed no difference when compared to WT mice in synapse type percentage (Figure 4G). The decreased Type II and increased Type III and upregulated I-B/I-A ratio were significantly restored in Lnx1^−/−; EphB2^F620D/F620D compound mutants compared with Lnx1^−/− mice (Figure 4H). Finally, to determine whether constitutive catalytic activation of EphB2 can rescue synaptic efficacy in Lnx1^−/− neurons, we measured synaptic activity in CA3 pyramidal neurons from acute brain slices of PW3 mice. Both frequency and amplitude of mEPSC were recovered to WT level in Lnx1^−/−; EphB2^F620D/F620D compound mutants (Figure 4I). Taken together, our data are consistent with a model whereby Lnx1 serves as a specific protein stabilizer for postsynaptic EphB receptors in hippocampal CA3 pyramidal neurons to promote diversiform spinogenesis and synaptic remodeling (Figure 5).