Calsyntenin-1 Negatively Regulates ICAM5 Accumulation in Postsynaptic Membrane and Influences Dendritic Spine Maturation in a Mouse Model of Fragile X Syndrome

INTRODUCTION Dendritic spines are small protrusions on the surface of neuronal dendrites that form the postsynaptic component of the excitatory synapse and play important roles in learning and memory. Alterations in dendritic spines are found in many types of mental retardation and other neurological disorders (Calabrese et al., 2006). By morphology, dendritic spines are usually classified as mushroom shaped, thin (or elongated), and stubby, and their shape is thought to correlate with the strength and activity of the synapse (Bourne and Harris, 2008). A mushroom spine has a bulbous head connected to the dendrite by a constricted neck (or stalk); thin spines have a smaller head, and stubby spines lack a neck. However, these categories are not separated by clear-cut boundaries but rather describe a continuum of shapes (Arellano et al., 2007). A distinctive feature of dendritic spines is the postsynaptic density (PSD), a large assembly of receptors and signaling proteins associated with the membrane of the spine head at the junction with a presynaptic bouton of the axon (Fifkova and Delay, 1982; Sheng and Hoogenraad, 2007). Dendritic spines are highly dynamic (Matus, 2005) and their formation, maturation, and plasticity heavily depend on the actin cytoskeleton remodeling (Ethell and Pasquale, 2005; Cingolani and Goda, 2008). However, the underlying mechanisms and even the structural organization of actin filaments in spines are poorly understood (Halpain, 2000; Rao and Craig, 2000; Ethell and Pasquale, 2005; Tada and Sheng, 2006), apparently because of their small size combined with complex organization. These features critically require electron microscopy (EM) to fully understand the biology of dendritic spines. The major types of actin filament organization across cell types and subcellular organelles include protrusive structures, such as branched networks in lamellipodia (Svitkina et al., 1997; Svitkina and Borisy, 1999) and parallel bundles in filopodia (Svitkina et al., 2003), and contractile structures, such as bundles and networks of linear filaments with mixed polarity (Verkhovsky et al., 1995). Early EM studies of dendritic spines by using thin-section (Fifkova and Delay, 1982; Markham and Fifkova, 1986) or freeze-fracture (Landis and Reese, 1983; Hirokawa, 1989) techniques detected long filaments, as well as a meshwork of short, potentially branched, actin filaments in spines, but they failed to provide a cohesive picture of their entire cytoskeleton. More recent light microscopic and functional approaches revealed that proteins normally involved in generation of protrusive branched networks, such as the Arp2/3 complex (Racz and Weinberg, 2008), WAVE1 (Kim et al., 2006; Hotulainen et al., 2009), cortactin (Hering and Sheng, 2003), N-WASP (Wegner et al., 2008), profilin (Ackermann and Matus, 2003), and cofilin (Racz and Weinberg, 2006; Hotulainen et al., 2009), were present in spines. However, myosin II (Cheng et al., 2000; Ryu et al., 2006) and α-actinin (Wyszynski et al., 1997) were also found there suggesting formation of contractile bundles or networks. Kinetically, dendritic spines contain dynamic and stable subpopulations of actin (Honkura et al., 2008), which may correspond to distinct types of actin filament arrays within spines. In general, a common belief is that the head of a spine should be similar to lamellipodia and contain an Arp2/3 complex-dependent branched network, whereas the spine neck is probably maintained by an axial actin filament bundle (Halpain, 2000; Rao and Craig, 2000; Tada and Sheng, 2006; Hotulainen et al., 2009). However, these ideas have not been directly proven and the actin filament organization in spines remains uncertain. Even less is known about the structure of presynaptic actin, which is believed to have a dual function of restraining synaptic vesicles and of directing them toward the synapse (Cingolani and Goda, 2008). Dendritic spines are thought to derive from dendritic filopodia that establish the initial contact with an axon, although other models also exist (Papa et al., 1995; Ethell and Pasquale, 2005; Yoshihara et al., 2009). The cytoskeletal organization and molecular composition of dendritic filopodia remain totally unknown. They may be similar to conventional filopodia found in neuronal growth cones and at the leading edge of other migrating cells that contain a tight bundle of long uniformly oriented actin filaments in their interior (Small et al., 2002; Svitkina et al., 2003; Korobova and Svitkina, 2008). Alternatively, dendritic filopodia may be structurally similar to dendritic spines (Papa et al., 1995), which structure as mentioned earlier is also unknown. These conflicting considerations require an experimental approach to investigate the actual structure of dendritic filopodia. Here, we used platinum replica EM to characterize the cytoskeletal organization and molecular composition of dendritic spines and dendritic filopodia in dissociated cultures of hippocampal neurons and revealed novel features of both structures essential for understanding of their morphogenesis and dynamics. MATERIALS AND METHODS Cell Culture and Transfection For high density cultures, hippocampal neurons were isolated as described previously (Wilcox et al., 1994) and were kindly provided by Dr. M. Dichter (The University of Pennsylvania, Philadelphia, PA). Every 5 d, one third of culture medium was replaced by fresh medium. For transfection with enhanced yellow fluorescent protein (EYFP)-actin (Clontech, Mountain View, CA) or mCherry-actin (Yang et al., 2009), hippocampal neurons were plated on 35-mm glass-bottomed dishes at 100,000 cells/dish concentration and transfected using Lipofectamine 2000 (Invitrogen, Carlsbad, CA) after 5–6 days in vitro (DIV). Transfected cells were analyzed on 14 DIV. Antibodies The following rabbit polyclonal antibodies were used: p16-Arc (Vignjevic et al., 2003), Arp3 (Santa Cruz Biotechnology, Santa Cruz, CA), and p34-Arc subunits of Arp2/3 complex (Millipore, Billerica, MA), nonmuscle myosin II from bovine spleen (Verkhovsky et al., 1987), and capping protein (from D. Schafer, University of Virginia). The following mouse monoclonal antibodies were used: fascin (Millipore), microtubule-associated protein (MAP2) (Sigma-Aldrich, St. Louis, MO), N-cadherin (Santa Cruz Biotechnology; gift from W. J. Nelson, Stanford University, Stanford, CA), α-tubulin (Sigma-Aldrich), and PSD-95 (Abcam, Cambridge, MA). Secondary fluorescently labeled antibodies and AlexaFluor 488-, 594-, and 647-labeled phalloidins were from Invitrogen, secondary rabbit or mouse antibodies conjugated to 12- or 18-nm colloidal gold were from Jackson ImmunoResearch Laboratories (West Grove, PA). Immunofluorescence and Light Microscopy Immunofluorescence staining of cells growing on glass coverslips was performed after cell extraction for 5 min with 1% Triton X-100 in PEM buffer [100 mM piperazine-N,N′-bis(2-ethanesulfonic acid)-KOH, pH 6.9, 1 mM MgCl2, and 1 mM EGTA] containing 2% polyethelene glycol (mol. wt. 35,000) and 5 μM phalloidin and fixation with 4% paraformaldehyde in phosphate-buffered saline (PBS) or 0.2% glutaraldehyde in 0.1 M Na-cacodylate, pH 7.3. Glutaraldehyde-fixed samples were quenched with 2 mg/ml NaBH4. For F-actin staining, fluorescently labeled phalloidin (0.033 μM) was added to the secondary antibody solution. For fascin immunostaining, extracted cells were fixed with 50% methanol in PBS and washed with PBS. For time-lapse imaging, cells were cultured in glass-bottomed dishes and kept on the microscope stage in the TC-MIS-20 × 46 miniature incubator (Bioscience Tools, Santa Clara, CA) at 36.5°C in the atmosphere containing 5% CO2 and 20% O2. For correlative immunostaining, marked coverslips were used to help with cell identification (Svitkina, 2007). Immediately after acquisition of 10- to 12-min-long movies with 2- to 3-s time intervals, the medium in the dish was changed to 4% paraformaldehyde in PBS, and cells were immunostained as described above. Light microscopy was performed using an inverted microscope (Eclipse TE2000; Nikon, Tokyo, Japan) equipped with Planapo 100 × 1.3 numerical aperture or 20 × 0.75 objectives and Cascade 512B charge-coupled device (CCD) camera (Roper Scientific, Trenton, NJ) driven by MetaMorph imaging software (Molecular Devices, Sunnyvale, CA). Electron Microscopy Samples for platinum replica EM, correlative light and electron microscopy, immuno-EM, and S1 decoration were processed as described previously (Svitkina, 2007; Korobova and Svitkina, 2008). For nonextracted samples, neuronal cultures were fixed sequentially with 2% glutaraldehyde in Na-cacodylate, and with aqueous solutions of 0.2% uranyl acetate, 0.1% tannic acid, and 1% OsO4. For Arp2/3 complex immunogold staining, cells were extracted as for immunofluorescence, fixed with 0.2% glutaraldehyde, quenched with NaBH4, and incubated in a mixture of p16-Arc, Arp3, and p34-Arc antibodies. For N-cadherin, myosin II, and capping protein immunostaining, unfixed cells extracted with 1% Triton X-100 and 4% polyethelene glycol in PEM buffer were incubated for 15 min with a primary antibody in the PEM buffer containing 5 μM unlabeled phalloidin, washed with the PEM buffer, fixed with 0.2% glutaraldehyde, stained with secondary antibody, and postfixed with 2% glutaraldehyde. For PSD-95 staining, cells were extracted/fixed for 5 min with a mixture of 0.5% Triton X-100 and 1% paraformaldehyde in PEM buffer with 5 μM phalloidin, washed in the same buffer, incubated with primary antibodies in PEM buffer for 15 min, postfixed with 2% glutaraldehyde, and stained with secondary antibody. Myosin S1 (gift from Dr. L. Tilney) was centrifuged at 100,000 × g for 30 min before use. For correlative light and electron microscopy, EYFP- or mCherry-actin–expressing cells were located on marked glass-bottomed dishes and phase-contrast and fluorescent images were acquired immediately before extraction and within 2 min after addition of the extraction solution. Then cells were fixed and processed for EM, as described previously (Svitkina, 2007; Korobova and Svitkina, 2008). Samples were analyzed using JEM 1011 transmission EM (JEOL USA, Peabody, MA) operated at 100 kV. Images were captured by ORIUS 835.10W CCD camera (Gatan, Warrendale, PA) and presented in inverted contrast. Identification of gold particles in replica EM samples was performed at high magnification after contrast enhancement to distinguish them from other bright objects in the samples. Thus, gold particles showed up as solid white circles in contrast to filament ends, for example, which usually have a donut-like appearance. Image Analysis and Statistics All morphometric measurements were done using MetaMorph or Photoshop (Adobe Systems, Mountain View, CA) software packages and repeated for at least two independent experiments. Data were analyzed using Excel software (Microsoft, Redmond, CA). Significance was determined using a two-tailed t test. For quantification of F-actin and Arp2/3 complex contents in filopodia, filopodia were manually selected on phalloidin-stained images using the MetaMorph selection tool and selection was transferred to corresponding Arp2/3 complex-stained images. Average fluorescence intensities of selected regions from both sets of images were recorded after background subtraction. To quantify the distribution of PSD-95 and N-cadherin immunogold labeling in dendritic spines relative to the adjacent axon, we measured the shortest distances between individual gold particles and the nearest microtubule or intermediate filament in the axonal shaft in EM samples. Gold particles occurring on the other side of the first axonal fiber were assigned negative numbers. Only gold particles located in the distal half of the spine, were included into quantification to exclude irrelevant labeling in the dendritic shafts where both PSD-95 and N-cadherin are also present. If the axon and the dendrite were not parallel, the distal half of the spine was determined by drawing a bisector for the angle between them until it intersects the spine. RESULTS Identification of Spines Platinum replica EM is advantageous for investigation of the cytoskeletal organization of whole-mount samples at high resolution (Svitkina, 2007). However, detergent extraction, which is required for this technique, dissolves membranes making cell boundaries undetectable and has a potential to perturb the cytoskeletal organization. Therefore, we performed control experiments to identify dendritic spines, determine their polarity, and test their preservation in EM samples. First, we performed replica EM of nonextracted hippocampal neurons in which cell boundaries remain preserved (Figure 1). Neurites in these cultures usually formed complex networks or aligned bundles (Figure 1A). Dendritic spines were abundant in mature cultures fixed after 14–28 DIV and could be recognized by their mushroom shape and/or ability to make contacts (presumably, synapses) with adjacent neurites (Figure 1, A–D). An axon interacting with the spine head was usually thinner than the dendrite, from which the spine emerged. The spine necks frequently expanded not only at the tip forming a bulbous head, but also at the root, although to a lesser extent, forming a delta-shaped base (Figure 1C). At 7–11 DIV, spines were rare, but dendritic filopodia abundant (Figure 1, E–H). Compared with straight and uniform growth cone filopodia in the same samples (data not shown), dendritic filopodia were very polymorphic. Occasionally, they made contacts with other neurites (Figure 1H). At this stage of development, dendrites also formed small lamellipodia-like protrusions (data not shown). Figure 1. Morphology of dendritic spines (A–D) and dendritic filopodia (E–H). Replica EM of nonextracted hippocampal neurons after 14 (A–D) or 10 (E–H) DIV. (A) Network of neurites with dendritic spines (arrows); two spines contact other neurites (axons). The cell body producing the major dendrite is located ∼20 μm away from the left margin of the panel. (B–D) Examples of mushroom (B), thin (C), and stubby (D) spines. (E) Distal region of a dendrite with polymorphic dendritic filopodia. (F–H) Examples of dendritic filopodia. Bars, 2 μm (A and E), 1 μm (B–D, F, and G), or 0.5 μm (H). We also stained hippocampal cultures with a dendritic marker MAP2 (Caceres et al., 1984) and phalloidin to distinguish dendrites and axons (Supplemental Figure S1). A strong MAP2 signal in dendrites correlated with prominent F-actin staining and a greater thickness of neurites; in contrast, MAP2-depleted axons contained much less F-actin and were on average thinner. Finally, we used correlative EM of EYFP-actin–expressing neurons to visualize the same spines sequentially by light and EM (Supplemental Figure S2). These results showed that detergent extraction and EM processing did not alter the shape or actin distribution in spines. They also revealed characteristic features of the spine cytoskeleton, such as a greater density of the actin filaments in spine heads, compared with spine bases. Based on these experiments, we applied following criteria to identify the spine polarity in EM samples (Supplemental Figure S3): a greater thickness of dendrites and abundance of actin filaments there, compared with axons; a greater size and higher actin filaments density in spine heads, as compared with bases; and a more distinct delta-like shape of bases, as compared with more polymorphic heads. Actin Cytoskeleton Organization in Dendritic Spines Mature 14–28 DIV hippocampal neurons were used for EM analysis of dendritic spines. Axons and dendrites could be recognized by axial bundles of microtubules and intermediate filaments in their interior, and dendrites also contained significant amount of actin filaments (Supplemental Figure S3). A typical mushroom spine contained three compartments: a bulbous head contacting the axon, a constricted neck in the middle, and a delta-shaped base at the junction with the dendrite (Figures 2 and 3 and Supplemental Figure S3). Figure 2. Cytoskeletal organization of dendritic spines: head and neck. EM of extracted 14 DIV neurons. Mushroom (A) and thin (B) spines associate with dendrites at the base (bottom) and with axons by the head (top). Unlabeled insets show the smaller versions of the corresponding main panels with axons color-coded in purple, dendrites in yellow, and spines in cyan. See Supplemental Figure S3 for determination of the orientation of the spine shown in A. Thick fibers in both neurites represent microtubules. Red asterisks indicate putative PSD fragments. Yellow boxes are enlarged in panels with corresponding numbers. Box 1, interaction of putative PSD (arrow) from the spine head with axonal intermediate filaments (green); actin filaments are shown in cyan and a microtubule in red. Boxes 2, 3, and 5, branched actin filaments (cyan) in the head (2), at the neck–head junction (3), and in the neck (5) of respective spines. Insets in panels 2 and 3 show nonpseudocolored regions outlined by yellow boxes. Dashed arrows in panel 3 indicate potential filament breakage. Box 4, interaction of actin filaments (cyan) with an axonal microtubule (red). Bars, 0.2 μm (A and B). Figure 3. Cytoskeletal organization of dendritic spines: base. EM of extracted neurons after 11 (A) or 14 (B and C) DIV shows undecorated (A and B) or S1-decorated (C) mushroom (A and C) or stubby (B) spines. The spine in A also may be considered stubby. Unlabeled insets show the smaller versions of the corresponding main panels with axons color-coded in purple, dendrites in yellow, and spines in cyan. Yellow boxes are enlarged in panels with corresponding numbers. Box 1, interaction of actin filaments (cyan) with axonal microtubules (red). Boxes 2 and 3, branched actin filaments (cyan) in the spine bases associate with dendritic microtubules (red). Box 4, linear actin filaments in the spine base have unbound ends (arrows) and are arranged in a delta-shaped configuration. S1-decorated actin filaments have a rough bumpy outline that distinguishes them from undecorated thin filaments (arrowheads). Bars, 0.2 μm (A–C). The spine head typically contained a dense network of short cross-linked actin filaments, in which branched filaments were clearly visible in relatively sparse regions (Figure 2A). Long filaments were also sometimes observed in the spine head. Surprisingly, the actin network in the dendritic spine head seemed to directly interact with axonal microtubules and intermediate filaments (Figure 2). Proximity of actin-rich spine heads to microtubules in the axon was also seen by fluorescence microscopy (Supplemental Figure S4). However, part of this network likely belongs to the presynaptic bouton, rather than to the dendritic spine (see below). Accordingly, occasional chunks of nonfibrillar material in the spine head, which might represent nonextracted PSD fragments, were found either next to the axonal microtubules and intermediate filaments, or slightly away from them (Figure 2A). In the spine neck, actin filaments formed an anisometric network, which consisted of loosely arranged actin filaments that might be longitudinally oriented, but only infrequently formed tight bundles. The number of filaments varied significantly along the length of the neck and many filaments did not span the entire neck. Filaments began or ended with unbound ends but also branched off of the side of another filament (Figure 2B). At the neck–head junction, the number of actin filaments abruptly increased toward the head, which usually occurred through extensive branching of neck filaments entering the head (Figure 2A). The spine base is a previously unrecognized structural compartment of the spine. It usually contained relatively long actin filaments converging from a broad area in the dendrite toward the neck in a delta-shaped configuration (Figures 2 and 3). These filaments either continued into the neck, or terminated within the base (Figure 3C). In many spines, the base also contained branched filaments (Figure 3A). Surprisingly, the actin filaments of the spine base frequently seemed to begin directly from the microtubule network in the dendrite and subsequently amplified through branching (Figures 2 and 3), whereas others originated from other actin structures in the dendrite. Some spines did not have a well-formed base (Figure 3B). Other morphological types of spines displayed the same structural plan. However, thin spines (Figure 2B) had longer necks and narrower heads, whereas stubby spines seemed to lack a neck and consisted of a patch of a densely branched actin network that likely represented the spine head or a merge of the base and the head (Figure 3B). The dimensions of spine compartments varied broadly even within one morphological class of spines blurring the distinction between the classes. Some mushroom-shaped dendritic protrusions not making contacts with axons had nonetheless a similar structural organization as the contact-forming spines (data not shown). Decoration of actin filaments with the myosin II subfragment 1 (S1) confirmed that actin filaments were the major cytoskeletal component of spines (Figure 3C). However, nondecorated filaments of ∼ 6 nm in thickness (including an ∼4-nm platinum layer) were also detected in different spine compartments (Figure 3C). Thus, the cytoskeletons of the head, neck, and base of a mature spine are not dramatically different from each other and consist of a mixture of branched and linear filaments, but in different combinations. Branched filaments dominate in the head, whereas linear filaments are more prominent in the base, and the neck may contain different ratios of both. The mutual alignment of actin filaments also varies between compartments being most prominent in the neck, but not to an extent of forming a tight bundle. This organization is conserved among all spine classes, mushroom, thin and stubby, but the dimensions of individual compartments vary broadly up to complete absence of a neck and/or a base. Molecular Markers in Dendritic Spines The presence of branched actin networks in different spine compartments suggested involvement of the Arp2/3 complex. Indeed, immunofluorescence staining detected Arp2/3 complex in the majority of heads and in 30–40% of necks and bases of mushroom or thin spines (Figure 4, A and D). In stubby spines, the Arp2/3 complex was found throughout the spine. Small actin patches that were usually present in dendrites also contained the Arp2/3 complex. Immunolocalization of another common marker of branched networks, heterodimeric capping protein (Svitkina et al., 2003), gave similar results (Figure 4E). An actin filament cross-linker, fascin, which is characteristic for conventional filopodia (Svitkina et al., 2003), was virtually absent from dendritic spines (Supplemental Figure S5). Figure 4. Immunofluorescence staining of cytoskeletal proteins in spines. (A–C) Low-magnification (top) and high-magnification (bottom) images of dendritic spines in 14 DIV hippocampal neurons stained with phalloidin (green) and indicated antibodies (red). Dendritic spines are marked by arrowheads. (D–F) Percentage of spine compartments immunopositive for indicated proteins in the population of spines. Data are collected from three experiments and expressed as mean ± SD (137 spines for the Arp2/3 complex; 90 spines for capping protein, and 121 spines for myosin II). Bars, 2 μm (A–C). Apparently seamless transition between dendritic and axonal cytoskeletons at the synaptic junction in the detergent-extracted samples prompted us to attempt to elucidate the position of a boundary between two cells using immuno-EM of N-cadherin, a transmembrane junctional molecule involved in synapse formation (Bourne and Harris, 2008), and PSD-95, a scaffolding protein in the PSD that has perijunctional localization (Tada and Sheng, 2006). As expected for the membrane-associated molecules, a significant fraction of both proteins was dissolved by the detergent treatment during immuno-EM processing, but a fraction of them remained. Although labeling was not dense, it was specific, as gold particles were not detected in inappropriate locations and they were absent when primary antibodies were omitted. Furthermore, the staining pattern was consistent between spines and experiments. Gold particles in both cases tended to localize at some distance from axonal microtubules and intermediate filaments within the actin network in spine “heads” (Figure 5A and Supplemental S6). Because the intensity of labeling was limited to approximately five to six gold particles per spine, it was not possible to precisely demarcate the interface between the axon and the spine in each individual case. Therefore, to quantitatively evaluate the label distribution and thus an average position of the boundary, we measured the distance between gold particles labeling N-cadherin or PSD-95 and the closest definite cytoskeletal component of an axon, usually a microtubule or an intermediate filament. The quantification produced similar results for both markers. Thus, gold particles labeling N-cadherin and PSD-95 were localized at distances of 264 ± 417 nm (mean ± SD; 10 spines, 59 gold particles) and 287 ± 382 nm (12 spines, 60 gold particles) from axons, respectively, supporting an idea that a fraction of the actin network at the spine tips, in the order of several hundreds nanometers, probably belongs to the axon rather than to the dendrite. The structure of the network was similar on the both sides of the putative boundary revealed by the N-cadherin or PSD-95 labeling. Additional gold particles of N-cadherin staining were also found in dendrites consistent with frequent bundling of dendrites in these cultures (Figure 1A). Figure 5. Immunogold EM of cytoskeleton-associated proteins in spines. Unlabeled insets show the smaller versions of the corresponding main panels with axons color-coded in purple, dendrites in yellow, and spines in cyan. (A) N-cadherin. (B) Arp2/3 complex. (C) Capping protein. (D) Myosin II. Gold particles showing protein localization are highlighted in orange. Red boxes are enlarged in panels with corresponding numbers to show noncolorized gold particles (arrowheads). Box 5 shows a linear cluster of gold particles at the bottom. Bar, 0.2 μm. Immunogold EM of Arp2/3 complex and capping protein gave similar results (Figure 5, B and C) and showed that all analyzed mushroom or thin spines (N = 23 and 22 spines for Arp2/3 complex and capping protein, respectively) contained gold particles in the head, and approximately half of spines was labeled in the neck (11 and 14 spines, respectively) and the base (13 spines for each protein). The density of gold particles in necks and bases was usually lower, than in heads. Stubby spines had even distribution of gold particles throughout the body. Next, we evaluated the precise distribution of myosin II in spines, which presence in spines was reported previously (Morales and Fifkova, 1989; Ryu et al., 2006). Immunofluorescence staining revealed a punctate, or sometimes diffuse, distribution of myosin II in dendritic spines, dendrites, and the cell body (Figure 4, C and F). Myosin puncta likely corresponded to myosin II bipolar filaments (Verkhovsky et al., 1995), whereas diffuse staining might correspond to individual cytoskeleton-associated myosin II molecules. In spines, myosin II puncta predominantly localized to the neck and a lower part of the head, and to a lesser extent to the base. Immunogold EM revealed similar distribution of myosin II between spine compartments (Figure 5D). Thus, of 11 mushroom or thin spines analyzed, the neck (10 spines) and the lower part of the head (9 spines) were most frequently labeled, whereas the base labeling was less frequent (5 spines). However, gold particles in the neck frequently formed linear clusters, possibly reflecting formation of myosin II filaments, whereas in the head they were usually scattered as single particles (Figure 5D). Actin Cytoskeleton Organization and Cytoskeletal Components in Dendritic Filopodia The dendritic filopodia were investigated by replica EM by using 10 DIV hippocampal neurons (Figure 6). Their structure was strikingly different from that of conventional filopodia (Svitkina et al., 2003; Korobova and Svitkina, 2008), but similar to that of spine necks. It consisted of loosely aligned actin filaments of varying length with some branched filaments and occasional bundles (Figure 6, A and B). In addition, patches of a branched network were associated with filopodial shafts and even more frequently with filopodial tips producing prominent bulges there (Figure 6, A and E). The base of dendritic filopodia was similar to that of spines, both in the shape and cytoskeletal organization (Figure 6). Decoration of actin filaments with myosin S1 revealed a small fraction of actin filaments with barbed ends facing away from the tip (Figure 6C), which is unusual for conventional filopodia. Besides filopodia, patches of highly branched actin network, either relatively flat like in lamellipodia (Figure 6G), or more three-dimensional (Figure 6F), were associated with dendrites in these cultures. Figure 6. Cytoskeletal organization of dendritic filopodia and patches. EM of extracted 10 DIV hippocampal neurons. (A) A typical filopodium containing actin network in the base and the shaft. (B) A rare type of filopodia with a relatively well-shaped actin filament bundle, which however contains branched filaments and linear filaments of variable lengths. (C) S1 decorated filopodium. Red boxes 1 and 2 are enlarged in numbered panels at right to show actin filaments of mixed polarity. Adjacent asymmetric units of S1 decoration are highlighted in alternating red and cyan colors. The compound orange arrowheads point toward the pointed ends of filaments. (D) Filopodium stained with myosin II antibody contains gold particles (orange) distributed along the length as linear clusters (boxes) or single particles. Red boxes 3 and 4 are enlarged in panels with corresponding numbers at left to show noncolorized gold particles (arrowheads). (E) Filopodium containing prominent patches of branched actin network at the tip and in the lower shaft (arrowheads). (F and G) Dendritic patches of branched actin network with three-dimensional (F) or flattened (G) morphology. Bars, 0.5 μm. By immunofluorescence staining, the Arp2/3 complex and capping protein were present in 65% (N = 54) and 69% (N = 74), respectively, of dendritic filopodia (Figure 7, A and B) unlike the conventional filopodia, which lack these proteins (Svitkina and Borisy, 1999; Svitkina et al., 2003). Conversely, fascin was absent in dendritic filopodia, but abundant in conventional growth cone filopodia (Figure 7D). A majority of dendritic filopodia (88%; N = 77) contained myosin II, especially in the proximal regions (80% of the myosin II-positive filopodia) (Figure 7C). Immunogold EM with the myosin II antibody revealed both single gold particles and their linear clusters in dendritic filopodia (Figure 6D). Figure 7. Molecular markers and dynamics of dendritic filopodia. (A–D) Fluorescence staining of cytoskeletal components in filopodia of 7–11 DIV hippocampal neurons by phalloidin (green) and indicated antibodies (red) shown at low magnification (top, A–C, left, D) or high magnification (bottom, A–C, right, D). Dendritic filopodia in A–C are indicated by arrowheads. Boxes 1 and 2 from D are enlarged in side panels with corresponding numbers. Unlabeled top panel shows a dendritic filopodium from another dendrite. Box 2 shows fascin-positive growth cone filopodia in contrast to dendritic filopodia not containing significant amounts of fascin. (E and F) Time-lapse phase-contrast sequences showing initiation of dendritic filopodia (arrows) from phase-dense spots (E) or small lamellipodia (F) in 10 DIV neurons. Bottom panel in E shows a color-coded image from the top panel to emphasize the phase-dense patches (purple). Time intervals are 6 s (E) or 3 s (F). Bars, 2 μm (A–C), 5 μm (F), and 5 μm (E and F). To understand how dendritic filopodia may be formed, we followed their dynamics in 10 DIV neuronal cultures by time-lapse phase-contrast microscopy. Filopodia usually originated from phase-dense dynamic patches or small lamellipodia associated with the dendritic shafts (Figure 7, E–F, and Supplemental Videos S1 and S2). These precursors of filopodia likely correspond to the patches of branched actin filaments we noted in EM samples (Figure 6F). Newly formed, as well as some preexisting filopodia, were highly dynamic, whereas other preexisting filopodia were stable and expressed relatively little dynamics. To address a possibility that stable and dynamic filopodia may differ in the contents of key cytoskeletal components, we evaluated the amounts of F-actin and the Arp2/3 complex by fluorescence microscopy in filopodia with pre-recorded behavior. Quantification of fluorescence intensities of filopodia stained with fluorescent phalloidin and Arp2/3 complex antibodies revealed no statistically significant differences in either F-actin or Arp2/3 contents between stable and dynamic filopodia (Supplemental Figure S7), consistent with our EM data showing comparable organization of the actin cytoskeleton in the entire population of filopodia. DISCUSSION Although the important role of dendritic spines in synaptic transmission is well established, the full understanding of their biology is critically delayed by the absence of their high-resolution structure. Diverse diagrams of the actin cytoskeleton organization in spines existing in the literature rather reflect a range of speculations than actual knowledge (Halpain, 2000; Rao and Craig, 2000; Ethell and Pasquale, 2005; Tada and Sheng, 2006), but they reveal how acutely this information is needed. In this study, we have characterized the cytoskeletal organization of dendritic spines and dendritic filopodia in their entirety and at high resolution and proposed a model of spine morphogenesis driven by actin cytoskeleton remodeling (Figure 8). Figure 8. Model for actin cytoskeleton organization in dendritic protrusions and for spine morphogenesis. The actin cytoskeleton in dendritic patches (left), dendritic filopodia (middle), and dendritic spines (right) has similar organization consisting of a mixed network of linear and branched actin filaments (blue) anchored to microtubules (red) or actin filaments in the dendritic shaft. The process of spine formation probably begins with the formation of a dendritic patch, which then elongates into a dendritic filopodium. On receiving of appropriate signals, the filopodium undergoes maturation into a spine; this process probably involves activation of the Arp2/3 complex followed by formation of a dense branched actin network, which drives the expansion of the filopodial tip into a spine head. Presynaptic bouton forming a synapse with the spine head spine contains a similar network of actin filaments. Membrane is shown in gray. Dendritic Spines Out of three structural domains of a mushroom spine, the base is a previously unrecognized compartment whose structure ranges from an elaborate mixture of branched and linear filaments to a few converging linear filaments. This variability resembles a situation with basal regions of conventional filopodia, in which fraction of linear filaments increases with filopodia maturation (Svitkina et al., 2003). This similarity suggests that the spine base may be a remainder of a precursor structure, which gradually changes its organization with its age. The spine neck, unexpectedly, is supported not by an axial bundle of actin filaments, but by a longitudinally stretched network of branched and linear filaments of different lengths, which are only roughly aligned with each other. Although we cannot completely exclude a possibility that this structure is an artifact of the technique, there are several arguments against such possibility. First, by correlative EM we do not reveal significant reorganization and/or loss of actin in spines. Second, the presence of Arp2/3 complex and capping protein in spine necks is also consistent with their network-like organization. Third, replica technique successfully reveals long bundled filaments in other cases (Svitkina et al., 2003; Yang et al., 2007; Korobova and Svitkina, 2008). Finally, our results do not actually conflict with previous EM data obtained by other techniques showing occasional episodes of bundling by thin section EM (Fifkova and Delay, 1982; Markham and Fifkova, 1986) or roughly aligned filaments by freeze-fracture EM (Landis and Reese, 1983; Hirokawa, 1989). Because detection of branched filaments and filament ends in sectioned or fractured samples is difficult, replica EM of whole-mount samples used here provides more interpretable images. The spine head undergoes constant actin-dependent shape changes (morphing), probably regulated by synaptic stimulation (Tada and Sheng, 2006; Bourne and Harris, 2008). Dynamic actin-dependent processes are frequently associated with branched actin networks nucleated by the Arp2/3 complex (Pollard and Borisy, 2003; Goley and Welch, 2006). Such networks were also expected to function in spine heads (see Introduction). Our data support these expectations by showing extensively branched actin network in the distal regions of spines. Although replica EM is very beneficial for revealing the cytoskeletal architecture, it is limited by necessity to remove the plasma membrane to expose the cytoskeleton. Consequently, it is not possible to determine precisely where the spine head ends. To overcome this problem, at least partially, we used immunogold labeling of N-cadherin and PSD-95 in conditions that partly preserve these plasma membrane-associated components. These data allowed us to suggest that an apparently single piece of actin network connecting a spine to the axon, in fact, consists of pre- and postsynaptic subsets. This idea is more consistent with the available data, compared with an alternative possibility of actin filaments from the spine head directly interacting with axonal microtubules. Indeed, transmembrane adhesion receptors mediating cell–cell junctions usually interact through additional proteins with the actin cytoskeleton on both sides of the junction, rather than with microtubules. Furthermore, there is convincing evidence that presynaptic actin exists and is important for synaptic vesicle trafficking (Cingolani and Goda, 2008). Our data suggest that the presynaptic actin cytoskeleton consists of a branched actin network similar to that in the spine head and is also associated with microtubules. In adaptive dynamic systems, like dendritic spines, protrusion and retraction cooperate in generating a proper shape, which may explain the presence of myosin II in spines (Morales and Fifkova, 1989; Ryu et al., 2006). We found that myosin II is biased toward the neck and proximal regions of the head, suggesting that contraction mainly occurs there. Filamentous myosin II in spines suggests the presence actin filaments of mixed polarity. However, we could not obtain enough filaments with interpretable polarity to test this prediction. Because the quality of decoration was good elsewhere, such as in growth cones, we assume that actin filaments in spines are overloaded with other actin-binding proteins preventing filament saturation with S1, as required for polarity determination. Dendritic Filopodia and Spine Morphogenesis Dendritic filopodia can transform into morphologically mature spines (Yoshihara et al., 2009). Their structure remained totally unknown but was assumed to resemble that of conventional filopodia. Strikingly, we found that dendritic filopodia had network-like cytoskeletal organization, which is unusual for highly elongated membrane protrusions, where a tight actin filament bundle is considered to be obligatory (Chhabra and Higgs, 2007). However, we have shown recently that bundled actin filaments are dispensable for maintaining filopodia induced by the membrane-deforming I-BAR domain of IRSp53, although polymerized actin is required (Yang et al., 2009). Dendritic filopodia provide a naturally occurring example of this kind. The network-like organization of dendritic filopodia probably makes them more plastic allowing for frequent changes of direction (Portera-Cailliau et al., 2003). The structural organization of dendritic filopodia suggests potential mechanisms of their differentiation into spines. By live imaging, the filopodium-to-spine transformation occurs as swelling of the filopodial tip and shortening (Marrs et al., 2001). The swelling may occur through Arp2/3 complex-dependent actin filament branching at the filopodial tip, which would drive the head expansion. Filopodia with sizable patches of branched actin network at the tip may represent transitional states in this process. The filopodia shortening during spine maturation was proposed to involve myosin II (Ryu et al., 2006). We indeed detected myosin II in the shafts of dendritic filopodia and also observed actin filaments of mixed polarity there. Both features are quite unusual for conventional filopodia, which are devoid of myosin II and have uniformly oriented filaments, but are consistent with contractile properties of dendritic filopodia. Another important question is the origin of dendritic filopodia. By light microscopy, they seem to grow directly from a dendrite (Dailey and Smith, 1996), usually from pre-existing actin patches (Andersen et al., 2005) or, in our experiments, from phase-dense spots or small lamellipodia. These sites of filopodia initiation likely correspond to patches of a branched actin network in dendrites, which may subsequently become the filopodial base, thus explaining the presence of branched filaments there. These branched filaments may be responsible for incorporation of labeled actin into roots of dendritic filopodia (Hotulainen et al., 2009). The mechanism of transformation of a relatively isometric patch into an elongated protrusion remains unclear. One possibility is contribution of membrane-deforming proteins, such as IRSp53 (Choi et al., 2005; Mattila and Lappalainen, 2008), which can induce and support tubular membrane protrusions (Saarikangas et al., 2009; Yang et al., 2009). Enhanced polymerization of actin filaments assisted by formin mDia2 may also contribute to this process (Hotulainen et al., 2009). Surprisingly, the actin structures in neurons frequently reside directly on the microtubule array. Moreover, some actin filaments seem to branch off of a microtubule in the neurite suggesting involvement of a microtubule-associated actin filament nucleator(s) or actin–microtubule cross-linkers, which remain to be identified. In conclusion, the cytoskeletal organization and molecular composition of dendritic spines and dendritic filopodia suggest a likely sequence of cytoskeletal reorganizations underlying the spine morphogenesis (Figure 8). We propose that the process begins with the formation of a small patch of branched actin network, which might be nucleated in association with dendritic microtubules. The patch subsequently elongates into a dendritic filopodium, which due to plastic network-like organization of its cytoskeleton is able to perform a wide range of movements searching for an axon. When an appropriate signal is received, it triggers the head formation by inducing extensive branching of actin filaments. Myosin II-dependent contractility within the head and the neck then modulates the shape of the spine to fit the requirements of synaptic transmission. This model provides a conceptual framework for future studies which would uncover structural reorganizations of the cytoskeleton occurring at different stages of synapse morphogenesis and specific roles of individual proteins in these processes. Supplementary Material [Supplemental Materials]

Fragile X syndrome is the most common form of inherited mental retardation. The disorder is mainly caused by the expansion of the trinucleotide sequence CGG located in the 5' UTR of the FMR1 gene on the X chromosome. The abnormal expansion of this triplet leads to hypermethylation and consequent silencing of the FMR1 gene. Thus, the absence of the encoded protein (FMRP) is the basis for the phenotype. FMRP is a selective RNA-binding protein that associates with polyribosomes and acts as a negative regulator of translation. FMRP appears to play an important role in synaptic plasticity by regulating the synthesis of proteins encoded by certain mRNAs localized in the dendrite. An advancing understanding of the pathophysiology of this disorder has led to promising strategies for pharmacologic interventions.

Dendritic spines are postsynaptic compartments of excitatory synapses that undergo dynamic changes during development, including rapid spinogenesis in early postnatal life and significant pruning during adolescence. Spine pruning defects have been implicated in developmental neurological disorders such as autism, yet much remains to be uncovered regarding its molecular mechanism. Here, we show that spine pruning and maturation in the mouse somatosensory cortex are coordinated via the cadherin/catenin cell adhesion complex and bidrectionally regulated by sensory experience. We further demonstrate that locally enhancing cadherin/catenin-dependent adhesion or photo-stimulating a contacting channelrhodopsin-expressing axon stabilized the manipulated spine and eliminated its neighbors, an effect requiring cadherin/catenin-dependent adhesion. Importantly, we show that differential cadherin/catenin-dependent adhesion between neighboring spines biased spine fate in vivo. These results suggest that activity-induced inter-spine competition for β-catenin provides specificity for concurrent spine maturation and elimination and thus is critical for the molecular control of spine pruning during neural circuit refinement.