The EphA4 receptor regulates dendritic spine remodeling by affecting β1-integrin signaling pathways

Dendritic spines are highly specialized microscopic protrusions on dendrites that typically have an enlarged head connected to the dendritic shaft by a narrow neck (Hering and Sheng, 2001; Ethell and Pasquale, 2005). Dendritic spines are the primary site where the postsynaptic components of excitatory synapses are located in the mammalian central nervous system. The spine neck serves to compartmentalize Ca²⁺ in the spine head away from the dendritic shaft, thus modulating excitatory synaptic transmission (Nimchinsky et al., 2002; Korkotian et al., 2004; Noguchi et al., 2005).

The formation and maintenance of dendritic spines require the coordinated activity of structural and signaling molecules (Ethell and Pasquale, 2005; Kennedy et al., 2005). Particularly important are proteins involved in the regulation of actin filaments, which are the main cytoskeletal component of spines that help define their shape (Matus, 2000). Time-lapse imaging has revealed that dendritic spines are dynamic structures that can undergo actin-based shape modifications and, in some cases, can form and disassemble within minutes (Matus, 2000; Hering and Sheng, 2001; Bonhoeffer and Yuste, 2002; Segal, 2005). Spine remodeling also occurs during the physiological stimulation of synapses and is correlated with changes in the strength of synaptic transmission. Structural reorganization of spines could be important for cognitive processes such as learning and memory (Yuste and Bonhoeffer, 2001; Nimchinsky et al., 2002; Segal, 2005). Indeed, several cognitive disorders are associated with spine malformations and changes in spine density (Fiala et al., 2002; Ethell and Pasquale, 2005). Thus, molecular dissection of the mechanisms involved in regulating dendritic spine morphology could be critical for understanding cognitive function and pathological conditions of the brain.

The Eph family of receptor tyrosine kinases and their ligands, the ephrins, play critical roles in both the developing and mature nervous system (Kullander and Klein, 2002; Poliakov et al., 2004; Yamaguchi and Pasquale, 2004; Pasquale, 2005). Eph receptors and ephrins interact at sites of cell–cell contact, where both are anchored to the cell surface. Ligand–receptor engagement elicits bidirectional signals: forward signals are generated through the Eph cytoplasmic region, including the tyrosine kinase domain, and reverse signals are communicated by the ephrin through cytoplasmic signaling molecules. Eph receptors and ephrins modify cell shape and movement by reorganizing the actin cytoskeleton. Several recent studies have shown that signaling by EphB receptors regulates dendritic spine morphogenesis, a process whereby the long and thin dendritic filopodial protrusions of immature neurons are replaced by dendritic spines in more mature neurons (Ethell et al., 2001; Henkemeyer et al., 2003; Penzes et al., 2003; Yamaguchi and Pasquale, 2004). Among the Eph receptors of the A class, EphA4 has been shown to regulate dendritic spine morphological plasticity in the adult hippocampus (Murai et al., 2003).

Signaling by EphA4, which is expressed on dendritic spines of pyramidal neurons, reduces spine length and density in acute hippocampal slices. A ligand for EphA4 in the hippocampus is ephrin-A3, which is glycosyl-phosphatidylinositol linked and localized on the surface of the astrocytic processes that surround spines. The molecular interplay between EphA4 and its ligand ephrin-A3 likely mediates a form of cell contact– dependent communication between astrocytes and neurons that helps to maintain the organization and architecture of dendritic spines by restricting their ability to grow and change shape (Murai et al., 2003). Indeed, EphA4 knockout mice have disorganized, longer, and more numerous spines than wild-type mice. Spine defects also accompany the introduction of a kinase-inactive form of EphA4 in pyramidal neurons, suggesting that kinase-dependent signals are involved in spine regulation. Therefore, identifying the signaling pathways that are activated downstream of EphA4 is important for understanding how dendritic spines can be structurally modified by cell surface receptors as well as for elucidating aspects of neuron–glia communication. In this study, we show that EphA4 regulates the morphological plasticity of dendritic spines by modulating integrin activity and downstream signaling molecules.

To investigate EphA4 signaling pathways involved in the regulation of dendritic spine morphology, we stimulated acute slices from postnatal mouse hippocampus with the ephrin-A3 ligand. Ephrin-A3 Fc, a soluble dimeric form of ephrin-A3, increased the tyrosine phosphorylation of EphA4 (Murai et al., 2003) and several other proteins (Fig. 1 a).

Src and Fyn are nonreceptor tyrosine kinases of the Src family that function downstream of EphA receptors and regulate the actin cytoskeleton (Ellis et al., 1996; Zisch et al., 1998; Bromann et al., 2004; Knoll and Drescher, 2004). Fyn in particular is present in mature synapses, plays a role in synaptic plasticity, is required for spatial learning and memory, and is needed for maintenance of dendritic spines (Grant et al., 1992; Morita et al., 2006). Interestingly, we found that an ∼120-kD tyrosine-phosphorylated protein (or proteins) coprecipitated with Fyn but not Src from hippocampal slices (Fig. 1 b), and ephrin-A3 Fc stimulation decreased the phosphotyrosine signal (Fig. 1 b). This suggests that one or more Fyn-associated proteins are dephosphorylated or become uncoupled from Fyn upon EphA receptor activation by ephrin in hippocampal slices. On the other hand, we did not detect changes in Fyn activation after ephrin-A3 Fc stimulation (Fig. 1 c).

To identify the tyrosine-phosphorylated proteins associated with Fyn, we examined the interaction of Fyn with ∼120-kD proteins known to bind Src family kinases, including the two closely related nonreceptor tyrosine kinases focal adhesion kinase (FAK) and proline-rich tyrosine kinase 2 (Pyk2), and the scaffolding protein Crk-associated substrate (Cas). Probing Fyn immunoprecipitates from hippocampal lysates for FAK or Pyk2 did not reveal a reproducible association (Fig. 2 a), although in some experiments, weak signals for FAK and Pyk2 were detectable (Fig. 2 a and not depicted). In contrast, substantial amounts of Cas coprecipitated with Fyn (Fig. 2 b). Both the SH2 and SH3 domains of Src family kinases can associate with the Src-binding domain of Cas (O'Neill et al., 2000; Bouton et al., 2001). Treatment of hippocampal slices with ephrin-A3 Fc decreased Cas binding to the SH2 domain of Fyn alone or fused with the SH3 domain in pull-down experiments (Fig. 2 c) and inhibited Cas association with full-length Fyn in coimmunoprecipitation experiments (Fig. 2 d).

As expected from the reduced Cas–Fyn association (Nakamoto et al., 1996; Ruest et al., 2001), ephrin-A3 Fc also decreased Cas tyrosine phosphorylation (Fig. 2 e). This inhibition of Cas phosphorylation was not observed in hippocampal slices from EphA4 knockout mice (Fig. 2 f and Fig. S2), suggesting that EphA4 is the main EphA receptor responsible for the effects of ephrin-A3 Fc in hippocampal slices. The reason for the lower basal levels of Cas phosphorylation in the EphA4 knockout slices remains to be determined.

Using a phosphospecific antibody, we found that ephrin-A3 Fc stimulation decreased phosphorylation of the substrate domain of Cas (Fig. 3 a), which contains multiple YxxP motifs. When phosphorylated, these motifs mediate binding of the SH2 domain of the adaptor protein Crk, which is an important effector of Cas (Feller, 2001; Chodniewicz and Klemke, 2004). Indeed, ephrin-A3 Fc treatment inhibited Cas binding to Crk in pull-down (Fig. 3 b) and coimmunoprecipitation experiments (Fig. 3 c). Phosphorylation of Crk on tyrosine 221 is also known to negatively regulate Crk interaction with Cas (Feller, 2001), but ephrin stimulation did not affect the phosphorylation of this site (Fig. 3 d). Thus, EphA4-induced dephosphorylation of Cas in the hippocampus disrupts Cas complexes.

Cas is a well-known regulator of cell shape and motility (O'Neill et al., 2000; Bouton et al., 2001) and, therefore, may play a role in dendritic spine regulation downstream of EphA4. Indeed, both EphA4 and Cas are present in brain synaptosomal preparations, including the postsynaptic density (PSD) fraction (Fig. 4 a). Furthermore, Cas expression in primary hippocampal neurons but not in glial cells (Fig. 4 b) supports the idea that Cas is involved in EphA4 forward signals that control dendritic spine morphology.

Biolistic transfection of a pool of four Cas siRNAs or individual Cas siRNAs in cultured hippocampal slices caused a substantial decrease in dendritic spine density (30–45%; Fig. 5) and a small but significant decrease in spine length (13–16%; P < 0.01) compared with transfection with control siRNAs that do not knock down Cas expression (Fig. 5 and Fig. S3). Cas siRNA transfection also decreased the number of mushroom-shaped spines, which represent the most common category of spines in hippocampal pyramidal neurons, and increased the number of stubby spines (Fig. 5). These effects were caused by the down-regulation of Cas expression because they were abolished when human Cas was cotransfected with a mouse-specific Cas siRNA (Fig. S4). Expression of the different Cas siRNAs did not detectably affect the overall morphology of the pyramidal neurons (unpublished data). These results implicate Cas as an important regulator of dendritic spines.

To identify the functional domains of Cas that play a role in dendritic spines, we transfected Cas deletion mutants (Fig. 6 a). Spine density was reduced by 48% in pyramidal neurons expressing the CasΔSB mutant (which lacks the Src-binding domain) and by 47% in neurons expressing the CasΔSH3 mutant (Fig. 6, a and b). In contrast, transfection of a Cas mutant that lacks the substrate domain (CasΔSD) and, therefore, cannot bind Crk did not significantly affect spine density (P > 0.05; Fig. 6, a and b). Spine length and width were not significantly affected by any of the Cas mutants (P > 0.05), with the exception of a small increase in spine width in the neurons transfected with CasΔSH3 and CasΔSB. With regard to spine shape, the CasΔSH3 and CasΔSB mutants caused a decrease in mushroom spines and a prominent increase in stubby spines, whereas CasΔSD decreased mushroom spines while increasing thin spines (Fig. 6 c). These results suggest that protein interactions mediated by the Cas SH3 and Src-binding domains contribute to maintaining spine density and play a role in defining spine shape.

The Cas SH3 domain is known to be a docking site for the FAK and Pyk2 kinases (O'Neill et al., 2000; Bouton et al., 2001), and we found that treatment of hippocampal slices with ephrin-A3 Fc decreases both FAK and Pyk2 tyrosine phosphorylation (Fig. 7, a and b). Because FAK, Pyk2, and Cas all become tyrosine phosphorylated upon integrin-mediated adhesion (Girault et al., 1999; Playford and Schaller, 2004), the coordinated decrease in their phosphorylation suggested that EphA4 activation may suppress integrin activity. Indeed, ephrin-A3 Fc stimulation of the hippocampal HT22 neuronal cell line, which expresses endogenous EphA4, caused a 35% decrease in cell attachment to surfaces coated with the integrin ligand fibronectin, whereas the integrin-independent attachment to poly-l-lysine was not affected (Fig. 7 c). Similar results were obtained with the neuronal-like B35 cell line, which stably expresses EphA4 (Fig. 7 d). Concurrent with the reduction in cell attachment, ephrin-A3 Fc stimulation reduced the level of Cas tyrosine phosphorylation in both cell lines (Fig. 7, c and d), which is similar to the effect observed in hippocampal slices.

Integrins are heterodimers containing an α and β subunit, and integrins containing the β1 subunit are highly expressed in the brain and influence hippocampal synaptic plasticity (Pinkstaff et al., 1999; Clegg et al., 2003; Chan et al., 2006; Z. Huang et al., 2006). To determine whether EphA4 may inactivate β1 integrins, we used the 9EG7 antibody, which, at low concentrations, detects the ligand-occupied (active) conformation of β1 integrins without interfering with the binding of extracellular matrix ligands (Bazzoni et al., 1995). Ephrin-A3 Fc treatment of HT22 cells caused a strong decrease in 9EG7 antibody binding (Fig. 7 e) without affecting the cell surface levels of β1 integrins, as determined by surface protein biotinylation (Fig. 7 f). Thus, EphA4 activation by ephrin-A3 inhibits β1-integrin activity and adhesion to fibronectin in neurons.

To examine whether integrins regulate dendritic spine morphology, we treated cultured hippocampal slices with an integrin function-blocking peptide (RGDFV) or a control peptide (RADFV; Fig. 8 a). Peptides containing the RGD (Arg-Gly-Asp) motif competitively antagonize integrin binding to extracellular matrix ligands (Ruoslahti, 1996; Shimaoka and Springer, 2003; Hama et al., 2004). Treatment of slices with the RGDFV peptide significantly decreased Cas tyrosine phosphorylation (P < 0.05; Fig. 8 b). Although RGD peptides have also been shown to promote integrin activation depending on the experimental conditions (Lin et al., 2003; Bernard-Trifilo et al., 2005; Shi and Ethell, 2006), this result is consistent with the expected inhibition of integrin downstream signaling pathways. The dendritic spines in slices treated with RGDFV peptide were 28% shorter than those treated with the control peptide (Fig. 8, a and c), whereas spine width was reduced only slightly, and spine density was not significantly altered (P > 0.05; Fig. 8 d). The shape of spines treated with the RGDFV peptide shifted from a mushroom-shaped to a stubby appearance (Fig. 8 e). These results show that integrin activity is required to maintain proper spine length and suggest that suppression of integrin function could play an important role in the effects of EphA4 on dendritic spine morphology.

To prevent β1-integrin inactivation during ephrin-A3 Fc stimulation, we treated acute hippocampal slices with high concentrations of the 9EG7 antibody. The antibody alone did not affect dendritic spine length, density, or width (Fig. 9, a–d; compare Fc with β1-Ab/Fc). As previously shown (Murai et al., 2003), treatment of hippocampal slices with ephrin-A3 Fc caused a significant decrease in spine length (P < 0.001; Fig. 9, a and b) and density (Fig. 9 c), with minor effects on spine width (Fig. 9 d, compare Fc with A3-Fc). In addition, as we show here, ephrin-A3 Fc decreased the proportion of mushroom-shaped spines while increasing stubby spines (Fig. 9 f). Remarkably, the 9EG7 antibody completely blocked all of the effects of ephrin-A3 Fc on the spines (Fig. 9, a–d; compare β1-Ab/Fc with β1-Ab/ephrin-A3 Fc). This is consistent with the lack of detectable Cas dephosphorylation when ephrin-A3 Fc was used together with the 9EG7 antibody (Fig. 9 e), although the antibody itself decreased Cas phosphorylation. We obtained very similar results when we treated hippocampal slices with the divalent cation Mn²⁺, which is known to promote the activation of many integrins, including β1 integrins (Fig. S5; Ivins et al., 2000; Arnaout et al., 2005). Thus, preventing β1-integrin inactivation counteracts the effects of EphA4 on dendritic spines.

This study provides the first direct evidence that EphA receptors regulate β1-integrin activity and downstream signaling pathways in neurons, which is crucial for governing the morphological plasticity of dendritic spines. Activation of EphA4 by ephrin-A3 may disrupt critical integrin-mediated attachment to the extracellular matrix and cell surface integrin ligands, thus inhibiting integrin signaling to Cas and other downstream signaling proteins. Inhibition of integrin function with an RGD antagonistic peptide and inhibition of Cas function with siRNAs decreased spine length and the proportion of mushroom-shaped spines while increasing stubby spines in hippocampal slices. These spine morphological changes are strikingly similar to those elicited by EphA4 activation with ephrin-A3 Fc, which is consistent with the idea that the different manipulations affect components of a common signaling pathway. However, the RGD peptide did not decrease spine density, suggesting that integrin inhibition by the peptide down-regulates Cas function less drastically than Cas siRNAs or that it may affect additional Cas-independent pathways.

Several studies have shown that Eph receptor forward signaling modulates the adhesion and migration of nonneuronal cells by affecting integrin function (Table S1 in Pasquale, 2005). Both positive and negative effects of Eph receptors on integrin-mediated adhesion have been reported, depending on the receptor involved and the cellular context. The interplay between Eph receptors and integrins may involve a direct physical association, as reported for EphA4 and αIIbβ3 integrin in platelets (Prevost et al., 2005).

Many integrin α and β subunits are expressed in the brain (Pinkstaff et al., 1999; Clegg et al., 2003). Among them, the β1 subunit, which can associate with various α subunits, is found in brain synaptosomal preparations and in the dendritic spines of cultured hippocampal neurons (Schuster et al., 2001; Clegg et al., 2003; Shi and Ethell, 2006). Signals mediated by integrins, including β1, have been shown to increase the length and density of dendritic protrusions and promote synapse formation and remodeling in cultured hippocampal neurons (Hama et al., 2004; Shi and Ethell, 2006). Furthermore, siRNA knockdown of the α5- or β1-integrin subunits in hippocampal neurons has been shown to decrease the number of dendritic protrusions and synapses (Webb et al., 2007; D.J. Webb, H. Zhang, and A.F. Horwitz. 2005. Society for Neuroscience. Abstr. 501.5). These findings are consistent with the idea that integrin inactivation contributes to the effects of EphA4 on dendritic spine morphology and density. The ability of a β1 integrin–activating antibody to block spine retraction downstream of EphA4 and the observed inhibition of β1-integrin activity by EphA4 suggest that integrins containing the β1 subunit play a particularly important role downstream of EphA4.

It is noteworthy that inhibiting integrin function with RGD peptides through the use of integrin function–blocking antibodies or by targeted gene deletion has been shown to cause defects in long-term potentiation (LTP), a form of synaptic plasticity (Chun et al., 2001; Chan et al., 2006; Z. Huang et al., 2006; Kramar et al., 2006). The effects of integrin inactivation on synaptic plasticity have been associated with decreased N-methyl-d-aspartate (NMDA) receptor function and defects in actin polymerization (Lin et al., 2003; Kramar et al., 2006). It will also be interesting to examine whether the changes in spine shape induced by integrin inactivation may have a functional impact on synaptic transmission and plasticity. Recent findings suggest that the induction of LTP, which is dependent on an increase in cytosolic Ca²⁺ concentration, may be more difficult to achieve in dendritic spines with thick neck regions such as stubby spines as a result of the faster dispersion of Ca²⁺ ions into the dendritic shaft (Nimchinsky et al., 2002; Korkotian et al., 2004; Noguchi et al., 2005). Inhibition of integrin activity may therefore moderate LTP induction and stabilization by EphA4 (Grunwald et al., 2004).

Consistent with a role of integrins in dendritic spine regulation and synaptic physiology, the extracellular matrix has also been shown to influence spine morphological plasticity. For example, mutant mice that have reduced levels of reelin, a large secreted glycoprotein that binds to the α3β1 integrin, have fewer and shorter spines compared with wild-type mice (Liu et al., 2001). Furthermore, degradation of extracellular matrix proteins by tissue plasminogen activator, which likely reduces integrin activity, has been implicated in the structural remodeling of spines that occurs in the visual cortex during experience-dependent plasticity (Mataga et al., 2004; Oray et al., 2004). Whether EphA4 may inhibit integrin activity by promoting proteolytic degradation of the extracellular matrix remains to be determined.

We have found that integrin inactivation by EphA4 results in the disassembly and inactivation of integrin signaling complexes, including the dissociation of Cas–Fyn complexes and the dephosphorylation of Cas, FAK, and Pyk2. The phosphatase mediating these dephosphorylation events remains unknown, although the tyrosine phosphatase SHP2 has been previously implicated in FAK dephosphorylation downstream of an EphA receptor in nonneuronal cells (Miao et al., 2000).

Even though Cas is most highly expressed in the brain and has been detected in complex with the NMDA receptor (Husi et al., 2000; J. Huang et al., 2006), its role in the nervous system is poorly characterized. Recent data have implicated Cas in axon elongation in cerebellar granule neurons (J. Huang et al., 2006), and our data show that Cas is preferentially expressed in hippocampal neurons rather than glial cells and is enriched in the PSD fraction of synaptosomal preparations. Furthermore, Cas knockdown in hippocampal slices decreased dendritic spine density and caused changes in spine morphology. The observed involvement of the Src-binding and SH3 domains of Cas in spine regulation further suggests that Cas may cooperate with Fyn and kinases of the FAK family, which interact with these two domains, to maintain proper spine morphology.

Interestingly, we detected Cas association with Fyn but not Src in hippocampal tissue. Although both Src and Fyn are components of the PSD and of NMDA receptor complexes (Salter and Kalia, 2004), differences in Src and Fyn protein interactions in the brain have been reported (Husi et al., 2000; Lauri et al., 2000). Thus, the synaptic functions of these two closely related kinases are not completely redundant. Accordingly, Fyn but not Src knockout mice demonstrate defects in the induction of LTP in adult CA1 hippocampal synapses (Grant et al., 1992; Kojima et al., 1997).

The dephosphorylation of FAK and Pyk2 that we observed in hippocampal slices stimulated with ephrin-A3 is consistent with reduced integrin function and suggests that inactivation of these two kinases may also contribute to spine morphological plasticity downstream of EphA4. However, the role of FAK in dendritic spines seems to be complex because FAK activation has been recently shown to be important for spine shortening and morphogenetic changes downstream of EphB receptors in cultured hippocampal neurons (Moeller et al., 2006).

Surprisingly, we did not detect significant changes (P > 0.05) in dendritic spines expressing the CasΔSD mutant, which does not bind Crk, except for a decrease in mushroom-shaped spines. Crk couples phosphorylated Cas to the guanine nucleotide exchange factor DOCK180, which activates Rac1, thus promoting actin polymerization and cell migration (Playford and Schaller, 2004). Therefore, either the Cas–Crk pathway does not play a major role in spine regulation or another signaling pathway can compensate for the disruption of Cas–Crk complexes. Indeed, EphA4 activation does not affect Crk phosphorylation on tyrosine 221, suggesting that the adaptor function of Crk is not impaired by EphA4 and that molecules other than Cas could still recruit Crk to promote Rac1 activation (Chen et al., 2004).

Our results demonstrate that EphA4 function in dendritic spines is closely tied to the regulation of β1-integrin signaling pathways, but additional pathways can also influence spine structure downstream of EphA4. For example, in neurons, EphA4 promotes the activity of at least two exchange factors that activate Rho family GTPases, Ephexin-1, and Vav2 (Shamah et al., 2001; Cowan et al., 2005; Sahin et al., 2005). Recently, a pathway involving the serine-threonine kinase Cdk5 and Ephexin-1 has been reported to decrease dendritic spine density downstream of EphA4 (Fu et al., 2007). This pathway likely affects dendritic spines through the activation of RhoA, a GTPase that promotes actomyosin contraction (Bonhoeffer and Yuste, 2002; Ethell and Pasquale, 2005). Nevertheless, β1-integrin inactivation appears to be a crucial event because preventing it counteracts all EphA4-dependent spine modifications. Further studies will be necessary to fully elucidate the Eph forward and ephrin reverse signaling pathways that modulate the morphological plasticity of excitatory synapses.

Monoclonal antibodies to phosphotyrosine (clone 4G10), Src, PSD95, and polyclonal antibody to Pyk2 were obtained from Upstate Biotechnology. Monoclonal antibody to FAK, Cas, Crk, and β1 integrin were purchased from BD Biosciences. The phosphospecific antibodies to Crk tyrosine 221, Cas tyrosine 165, and Src tyrosine 416 were obtained from Cell Signaling. The monoclonal 9EG7 antibody to activated β1 integrin was obtained from BD Biosciences. Monoclonal antibodies to β-actin and syntaxin were purchased from Sigma-Aldrich. Antibodies to Fyn and SHP2 and a Cas polyclonal antibody were purchased from Santa Cruz Biotechnology, Inc. The monoclonal antibody to EphA4 was obtained from Zymed Laboratories.

Tissues or cells were lysed in Hepes buffer (50 mM Hepes, pH 7.5, 150 mM NaCl, 5 mM KCl, 1 mM EDTA, 10% glycerol, and 1% Triton X-100) or modified radioimmunoprecipitation assay buffer with 1 mM sodium orthovanadate, 1 mM NaF, 1 mM PMSF, and other protease inhibitors (Sigma-Aldrich). GST-Fyn SH2 and GST-Crk SH2 fusion proteins have been previously described (Zisch et al., 1998, 2000). Recombinant GST fusion proteins were affinity purified from Escherichia coli BL21 (Pfizer) by adsorption to glutathione–Sepharose 4B beads (Pfizer). GST-Fyn SH3/SH2 fusion protein was purchased from Santa Cruz Biotechnology, Inc. For GST pull-down assays, cell lysates were incubated with 10 μg GST fusion proteins immobilized on glutathione beads. For immunoprecipitations, cell lysates were incubated with 2–5 μg of antibodies and GammaBind Plus Sepharose beads (GE Healthcare) for 3 h at 4°C. Protein extracts were separated by SDS-PAGE and probed by immunoblotting. Detection of HRP-conjugated secondary antibodies (GE Healthcare) was performed with enhanced chemiluminescence detection systems (GE Healthcare or Pierce Chemical Co.).

All steps were performed at 4°C as described previously (Rogers and Lemaire, 1991). Freshly dissected adult mouse brains were homogenized in 10 vol of ice-cold homogenization buffer (buffer H [50 mM Tris-HCl, pH 7.4, 1 mM CaCl₂, 1 mM MgCl₂, and 1 mM NaHCO₃] containing 320 mM sucrose) using a motor-driven glass-fluorinated ethylene polymer homogenizer. The homogenate was centrifuged at 700 g for 10 min to produce supernatant S1. S1 was further centrifuged at 17,000 g for 15 min to produce pellet P2. P2 was resuspended in homogenization buffer, layered on top of sucrose gradient (equal volumes of 1.2, 1.0, 0.8, and 0.65 M sucrose in buffer H), and centrifuged at 100,000 g for 2 h. A turbid fraction was recovered from the 1.0 /1.2 M sucrose interface and resuspended in at least 2 vol of homogenization buffer. The suspension was centrifuged for 30 min at 100,000 g to produce the pellet P2′ (synaptosome fraction). P2′ was resuspended in Tris buffer (50 mM Tris-HCl, pH 7.4, and 1 mM EDTA with protease inhibitors), incubated for 45 min on ice, and centrifuged at 100,000 g for 30 min. The supernatant containing soluble synaptic proteins was collected (synaptic cytosol fraction). The membrane pellet was resuspended in homogenization buffer, applied on top of a five-step discontinuous sucrose gradient (1.2, 1.0, 0.8, and 0.65 M sucrose), and centrifuged at 100,000 g for 2 h. Pure synaptic membranes were recovered from the 1.2/1.0 M sucrose interface and diluted with at least 2 vol of homogenization buffer (synaptic membrane fraction). A pure membrane fraction was pelleted at 100,000 g for 30 min, resuspended in homogenization buffer, and mixed with an equal volume of 3% Triton X-100 in homogenization buffer. Membranes were incubated for 30 min on ice and centrifuged at 100,000 g for 2 h to produce a supernatant (soluble membrane fraction) and a Triton X-100–insoluble material pellet. Triton X-100–insoluble proteins were resuspended in homogenization buffer containing 1.5% Triton X-100 and applied on top of a 0.83-M sucrose cushion. After centrifugation at 105,000 g for 1 h, PSDs were collected as the resulting pellet and resuspended in Tris buffer containing 0.25% Triton X-100 (PSD fraction). Equal amounts of protein from synaptosome, synaptic cytosol, synaptic membrane, soluble membrane, and PSD fractions were analyzed by immunoblotting.

Primary hippocampal neurons were prepared from embryonic day 17 rat embryos and cultured as described previously (Zafra et al., 1990) with minor modifications. In brief, after trypsinization and mechanical dissociation, hippocampal cells were suspended in DME supplemented with 10% FCS and preattached on uncoated culture plates for 2 h to remove glial cells. Neurons recovered as nonadherent cells were plated on poly-d-lysine– (50 μg/ml) and laminin (5 μg/ml)-coated plates at a density of 3 × 10⁴ cells/cm² in neurobasal medium supplemented with B27, l-glutamine, and penicillin/streptomycin. Neurons were cultured for 14 d in the presence of 2 μM AraC to prevent the growth of contaminating glial cells. The glial cells separated from the neurons by attachment on uncoated culture plates were cultured for 14 d in DME supplemented with 10% FCS, l-glutamine, and penicillin/streptomycin.

Hippocampal slices (300 μm in thickness) were prepared and stimulated for 45 min as previously described using the middle one third of the hippocampus (Stoppini et al., 1991; Murai et al., 2003). Slices were kept for 2–5 min in MEM (Invitrogen) supplemented with 25 or 5% horse serum (Invitrogen), 25% HBSS (Irvine Scientific), and 0.65% dextrose before stimulation. In some experiments, 9.5 μg/ml ephrin-A3 Fc (R&D Systems) was preclustered with 1 μg/ml anti-Fc antibodies, which produced similar EphA4 phosphorylation and Cas dephosphorylation as unclustered ephrin-A3 Fc. To promote β1-integrin activation, slices were pretreated with 20 μg/ml 9EG7 antibody for 25 min or 500 μM Mn²⁺ for 20 min before stimulation with Fc proteins. Slices were either lysed and analyzed by immunoblotting or fixed in 4% PFA in 0.1 M PBS for 30 min, labeled with DiI crystals (Invitrogen), and analyzed by confocal z-series imaging as previously described (Murai et al., 2003). EphA4 knockout mice were provided by A. Boyd and M. Dottori (Queensland Institute of Medical Research, Brisbane, Australia).

A gene gun (Helios; Bio-Rad Laboratories) was used to transfect plasmids into pyramidal neurons. Hippocampal slices prepared as described in the previous paragraph were cultured for 6 d on 0.4-μm Millicell-CM organotypic inserts (Millipore) in MEM containing 25% horse serum, 25% HBSS, and 0.65% dextrose. Slices were then transfected (80 pounds per square inch) with pSSRα/farnesylated enhanced GFP (EGFP-F; control), Cas/EGFP-F, CasΔSD/EGFP-F, CasΔSH3/EGFP-F, or CasΔSB/EGFP-F plasmids ((25 μg Cas plasmid mixed with 20 μg EGFP-F plasmid coated onto 25 mg of 1.6-μm gold particles; Nakamoto et al., 1996). Cas plasmids were provided by K. Vuori and A. Motoyama (Burnham Institute for Medical Research, La Jolla, CA). After 48 h, the slices were fixed in 4% PFA in 0.1 M PBS for 30 min before z-series imaging of dendrites from transfected CA1 pyramidal neurons. For integrin inhibition, 170 μM RGDFV or RADFV (control) peptide (BIOMOL Research Laboratories, Inc.) was added to the media of EGFP-F–transfected cultured hippocampal slices. After an additional 24 h in culture, the slices were analyzed either by confocal z-series imaging or immunoblotting.

Four siRNAs for mouse Cas were obtained from Dharmacon, including siCas #1 (GCATAGGGCATGACATCTA; nucleotides 678–696; GenBank/EMBL/DDBJ accession no. NM_009954), siCas #2 (AGACGGAGCAGGATGAGTA; nucleotides 804–822), siCas #3 (CCGAGGAAGTCCAGGATTC, nucleotides 1,816–1,834), and siCas #4 (GATTCTGGTTGGCATGTA, nucleotides 292–310). Two scrambled siRNAs (nontargeting siRNA #1 and #2; Dharmacon) were used as controls. To obtain the pEGFP-hCas vector, human Cas cDNA (GenBank/EMBL/DDBJ accession no. BC062556; Open Biosystems) was amplified by PCR with the forward primer 5′-GCTAGCCTCGAGCATGAACCACCTGAACGT-3′ and the reverse primer 5′-GCTAGCGAATTCTCAGGCGGCTGCCAGACA-3′. The PCR product was digested and inserted into the XhoI and EcoRI sites of pEGFP-C2 (BD Biosciences) and verified by sequencing.

Hippocampal slices were cotransfected with the siRNAs and the indicated plasmid using a gene gun as described previously (Boda et al., 2004; Govek et al., 2004). 24 μg of duplex siRNA and 12 μg of plasmid were mixed and coated onto 10 mg of 1.6-μm gold particles. Transfected (green fluorescent) dendrites were analyzed 2 d after transfection by confocal z-series imaging. The effectiveness of siRNA transfection into hippocampal pyramidal neurons was verified by using a rhodamine-labeled siRNA transfected together with EGFP-F. The effectiveness of Cas knockdown by the siRNAs and the rescue of Cas expression by EGFP-hCas was verified in NIH3T3 cells. Cells were plated at 80% confluency, cultured overnight in DME and 10% bovine calf serum, and transfected in Opti-MEM with 78 nM siRNAs (with or without plasmids) using LipofectAMINE 2000 (Invitrogen). After 4 h, the medium was replaced by the DME culture medium, and Cas expression was assessed after 48 h by immunoblotting of cell lysates.

Images were collected using a confocal microscope (MRC 1024; Bio-Rad Laboratories) and analyzed using ImageJ 1.62 software (National Institutes of Health [NIH]), and the analysis of spines was confined to secondary and tertiary dendrites from the stratum radiatum. Quantification of spine lengths and widths was performed as described previously (Murai et al., 2003). In brief, the length was measured from the tip of spine head to the interface with the dendritic shaft. The width was taken as a diameter of the spine head perpendicular to the length of the spine. Spines were assigned to morphological categories as described previously (Maruoka et al., 2005) by an investigator blind to the conditions. Mushroom spines are <2 μm in length, >0.5 μm in width, and are connected to the dendritic shaft by a narrower portion (neck). Stubby spines are <2 μm in length, >0.5 μm in width, and lack a defined neck. Thin spines are <2 μm in length, <0.5 μm in width, and have a neck. Filopodia spines are >2 μm in length, <0.5 μm in width, and do not have a distinct spine head. Irregular spines are branched spines (two heads and two necks merged into a single neck at the base) and multilobed spines (many heads fused together on top of a single neck). All measurements were obtained by counting 478–2,182 spines per condition (from 6–13 neurons) from independent experiments. Spine cumulative distributions were compared using the Kolmogorov-Smirnov test, and means were compared using the t test or analysis of variance (ANOVA) test followed by Dunnett's, Tukey's, or Bonferroni's posthoc tests.

To obtain the EphA4-B35 cells, the rat B35 neuronal-like cell line (Schubert et al., 1974) was transfected with the EphA4 cDNA in pcDNA3 using SuperFect (QIAGEN) and selected for 2 wk with 500 μg/ml G418. To perform the cell attachment assays, the HT22 hippocampal neuron cell line (Li et al., 1997) or EphA4-B35 cells were starved overnight in DME with 1% BSA, detached, and kept in suspension for 30 min at 37°C in DME–1% BSA. Cells (7 × 10⁴ cells per well of a 12-well plate) were then allowed to attach to poly-l-lysine– (20 μg/ml; Sigma-Aldrich) or fibronectin-coated coverslips (20 μg/ml fibronectin for EphA4-B35 and 10 μg/ml for HT22; Chemicon) for 10 min at 37°C in the presence of 10 μg/ml Fc or 10 μg/ml ephrin-A3 Fc. Attached cells were fixed in 4% PFA, stained with DAPI (Invitrogen), and counted blind (seven to nine 20× microscope fields were counted per condition). To determine Cas phosphorylation, both adherent and nonadherent cells were pooled, lysed, and probed by immunoblotting. HT22 and B35 cell lines were provided by W. Stallcup and B. Moosmann (Burnham Institute for Medical Research, La Jolla, CA), respectively.

HT22 was starved and plated on tissue culture plates coated with 10 μg/ml fibronectin for 1 h at 37°C in the presence of 10 μg/ml Fc, 10 μg/ml ephrin-A3 Fc, or 1 mM MnCl₂. To determine the levels of activated β1 integrin, live cells were rinsed with PBS and incubated with 2 μg/ml 9EG7 rat monoclonal antibody or control rat IgG for 15 min at 37°C. Cells were rinsed with PBS and lysed in SDS sample buffer, and bound antibodies were detected by immunoblotting with biotin-conjugated anti–rat IgG antibody (Vector Laboratories) followed by HRP-streptavidin (Zymed Laboratories). To determine the levels of cell surface β1 integrin, cells were washed with PBS, and surface proteins were labeled with 1 mM sulfosuccinimidyl-6-(biotin-amido)hexanoate (Pierce Chemical Co.) for 30 min at 4°C. Cells were washed with PBS and lysed in Hepes buffer with protease inhibitors, and biotin-labeled proteins were incubated with 50 μl streptavidin-conjugated beads (Biochemika) for 1 h at 4°C. Bound proteins were eluted and analyzed by immunoblotting with anti–β1-integrin antibodies.

Protocols for the animal studies were approved by the Burnham Institute's Animal Research Committee.

Fig. S1 summarizes the quantification and statistical analysis of the data from immunoblotting experiments. Fig. S2 confirms that ephrin-A3 Fc decreases Cas tyrosine phosphorylation in hippocampal slices from EphA4^+/− mice but not in slices from EphA4^−/− mice. Fig. S3 confirms that Cas knockdown decreases dendritic spine density and length in hippocampal slices by using a second siRNA control. Fig. S4 shows that the expression of human Cas abrogates the effects of mouse Cas siRNA on dendritic spine morphology. Fig. S5 demonstrates that integrin activation with Mn²⁺ blocks ephrin-A3 Fc–induced spine changes.

We thank F. Valencia for excellent technical assistance, S. Codeluppi for help with biolistic transfections, K. Vuori and A. Motoyama for Cas plasmids, A. Boyd and M. Dottori for the EphA4 knockout mice, W. Stallcup for the B35 cell line, and B. Moosmann for the HT22 cell line.

This work was supported by NIH grants to E.B. Pasquale as well as postdoctoral fellowships from the Fondation pour la Recherche Médicale to C. Bourgin, from the NIH to K.K. Murai, and from the Deutsche Forschungsgemeinschaft to M. Richter.