• We are sorry, but NCBI web applications do not support your browser and may not function properly. More information
Logo of pnasPNASInfo for AuthorsSubscriptionsAboutThis Article
Proc Natl Acad Sci U S A. Aug 26, 2008; 105(34): 12587–12592.
Published online Aug 22, 2008. doi:  10.1073/pnas.0711669105
PMCID: PMC2519044
Neuroscience

Structural plasticity with preserved topology in the postsynaptic protein network

Abstract

The size, shape, and molecular arrangement of the postsynaptic density (PSD) determine the function of excitatory synapses in the brain. Here, we directly measured the internal dynamics of scaffold proteins within single living PSDs, focusing on the principal scaffold protein PSD-95. We found that individual PSDs undergo rapid, continuous changes in morphology driven by the actin cytoskeleton and regulated by synaptic activity. This structural plasticity is accompanied by rapid fluctuations in internal scaffold density over submicron distances. Using targeted photobleaching and photoactivation of PSD subregions, we show that PSD-95 is nearly immobile within the PSD, and PSD subdomains can be maintained over long periods. We propose a flexible matrix model of the PSD based on stable molecular positioning of PSD-95 scaffolds.

Keywords: dendritic spine, glutamate, postsynaptic density, synapse, PDZ scaffold

The postsynaptic density (PSD) at excitatory synapses in the brain is a proteinaceous ‘organelle’ that positions neurotransmitter receptors across the synaptic cleft from sites of neurotransmitter release and links postsynaptic receptors with intracellular signaling cascades. The dynamic regulation of individual PSDs forms the basis for current molecular theories of learning and memory (13). Determining the internal architecture that controls PSD organization is particularly important because the size and shape of individual PSDs correlates with synapse function. In particular, larger and more complex PSDs are associated with more postsynaptic receptors and are apposed to axon terminals with larger active zones and more synaptic vesicles (47). In addition, average PSD size and morphological complexity vary over development (8) and are altered in disease (9). Such data have been interpreted to indicate that individual PSDs are dynamic structures that change shape (10, 11). However, the extent to which individual PSDs are structurally plastic, and the basis for variation in PSD size, shape, and organization remains unknown.

Proteomic characterization of the PSD (8, 1214), the atomic structure of many PSD proteins (15), the 3-dimensional shape of biochemically extracted PSDs (16), and the steady-state position of molecules within the PSD (1618) have been determined to increasing accuracy. Recent live-imaging experiments have focused on the accumulation and dispersal of PSD scaffolds during synaptogenesis or synapse loss (1922), the translocation of assembled PSD scaffolds in developing neurons as synapses form (23), or the rates of exchange of various PSD constituents (22, 2426). However, the submicron size of the PSD has hindered analysis of its morphology and internal architecture in living neurons. In particular, it remains unknown whether individual PSDs at established synapses undergo changes in size and shape and how PSD structural plasticity relates to fluctuations in PSD molecular spacing or content. Furthermore, whether scaffold proteins move while in the PSD remains untested. In the present study, we monitored internal PSD structural plasticity and the movement of scaffold proteins within the PSD on submicron scales using high-resolution live cell imaging.

Results

PSDs Are Structurally Dynamic at Established Synapses.

We performed time-lapse morphometric analysis of hippocampal neurons expressing GFP-tagged PSD-95, an abundant synaptic PDZ protein that delineates the PSD of excitatory synapses (15, 16). The PSD occupied only a fraction of the spine (Fig. 1A), and most PSDs in identified, non-overlapping spines were much larger than diffraction-limited objects (Fig. 1B), allowing detailed analysis of their shape. All PSDs persisted for the duration of our imaging experiments, but individual PSDs underwent extensive morphological alterations on the time scale of minutes [Fig. 1C and supporting information (SI) Movie S1]. By co-expressing PSD-95 with other proteins of the PSD, we found that identical changes occurred simultaneously for multiple PSD proteins. For example, stargazin-GFP (Fig. 1D and Movie S2), Shank-GFP (Movie S3), and GKAP1-GFP (data not shown) faithfully colocalized with PSD-95-mCherry, and the protein pairs displayed rapid and essentially identical reshaping. Thus, reshaping of PSD-95-GFP reflects architectural rearrangement of the large ensemble of PSD components.

Fig. 1.
Individual PSDs undergo continuous morphological plasticity. (A) Individual spines from hippocampal neurons 4 weeks in culture expressing PSD-95-GFP. Scale bar, 1 μm. (B) Distribution of PSD sizes in spines of hippocampal neurons 4 weeks in culture, ...

To quantify ongoing PSD structural change, we first examined its time course by acquiring pairs of high-speed image z-stacks (lasting ≈300 ms per stack) at varying intervals. We subtracted the paired images and totalled the intensity difference resulting from reshaping of PSDs within regions encompassing the synapse (Fig. 2A). At very short intervals (< 0.5 sec), little structural change could have taken place, so the difference value depends only the imaging noise. At long intervals (64 sec), difference scores were consistently large, indicating a change in shape. The intervening intervals revealed an increasing degree of movement discernible at times as small as 4 sec and fit by a curve with a half-time of just 16 sec. Thus, individual PSDs are subject to rapid structural plasticity on the time scale of seconds.

Fig. 2.
PSD structural plasticity is rapid, graded, continuous, and regulated by synaptic activity. (A) Rapid time course of PSD change. Pairwise subtraction of images was used to measure intensity differences at individual PSDs (n = 220 PSDs in 5 cells) over ...

To investigate the nature of PSD structure change in more detail, we performed morphometric analysis of time lapse images. Among a variety of parameters tested, we focused our analysis on PSD elliptical form (EF = length/breadth, Fig. 2B), a sensitive measure of PSD shape that, unlike PSD area or perimeter, was not appreciably affected by thresholding procedures during analysis or by photobleaching during long time-lapses. This quantification revealed that spontaneous changes in PSD structure occurred in a graded fashion (Fig. 2B and Movie S4), and were not abrupt alterations in form as would be expected if PSD components were added or subtracted en bloc. In addition, changes in PSD shape were not accompanied by changes in fluorescence intensity (Fig. 2B, Lower) indicating that the number of PSD-95 molecules remained constant during PSD structural plasticity. Further, we found that PSD dynamics at neighboring synaptic contacts were asynchronous (Fig. 2C and Movie S5), indicating that the regulation of PSD structure is local and synapse-specific. Despite large fluctuations of the morphology of individual PSDs, the population of PSDs showed a stable average EF (Fig. 2D). Thus, individual PSDs at established synapses are structurally dynamic, and this is not a directed or maturational evolution of the population related to synapse growth or loss. Rather, ongoing changes in PSD shape are an intrinsic feature of otherwise stable synapses.

We noted that individual PSDs entered periods of relatively high dynamic behavior. We delineated the periods when the rate of morphological change was high (Mode 1) or low (Mode 0) as defined by periods where the absolute value of the instantaneous rate of change of the EF over time was either greater or less than the standard deviation (Mode 1, |dEF/dt| ≥ σ; Mode 0, |dEF/dt| < σ; Fig. 2E). We found that 87 ± 3% of PSDs entered Mode 1 at least once within 25- to 60-min recordings, and the average duration of occupancy in Mode 1 was 3.4 ± 0.2 min. Although individual PSDs entered and exited this mode periodically, the overall number of PSDs in Mode 1 (27 ± 2%) remained stable for long periods (Fig. 2F) after an initial drop which we attribute to accommodation to the buffer and temperature of the imaging conditions. Notably, 57 ± 4% of the total change in EF occurred during Mode 1, despite accounting for only 27% of the total time examined. Thus, PSD structural plasticity is maintained by frequent entry into a state of rapid morphological alteration.

To test whether synaptic activity regulates PSD structural dynamics, we first reduced global activity in our cultures by chronically inhibiting action potentials with tetrodotoxin (TTX). As an index of the degree of morphological change at PSDs, we measured the coefficient of variation (CV) of PSD EF. Chronic TTX treatment did not affect this index (Fig. S1A), indicating that PSD dynamics are maintained following homeostatic increase of synapse strength (27). To test the response to acutely increased activity, we restored activity in chronically blocked cultures with an abrupt switch to solution containing the GABAA receptor antagonist bicuculline (Bic) and the potassium channel blocker 4-aminopyridine (4-AP). This treatment did not change the average PSD EF (Fig. S1B) but prompted a pronounced and transient increase in the CV of EF (Fig. 2 G and I). This transient increase was prevented by including the ionotropic glutamate receptor antagonists DNQX and APV in the activity-promoting solution (Fig. 2 H and I), suggesting that the response is driven by excitatory synaptic activity. Intriguingly, neurons treated with Bic/4-AP experienced an abrupt spike in the fraction of PSDs occupying the dynamic Mode 1, compared to control neurons treated also with DNQX/APV (Fig. 2J). Thus, structural dynamics of the PSD are acutely regulated by synaptic activity. Together, these findings suggest that increased alterations of PSD form are part of a rapid remodeling process synapses undergo during accommodation to changes in network activity. It will be important to learn how control of PSD-95 scaffold dynamics relates to the reported regulation of PSD-95 protein accumulation, dispersal, and degradation at synapses (13, 21, 23, 24, 2830).

Shifting PSD-95 Scaffold Density Within Single PSDs.

Changes in PSD size and shape may reflect the local scaffold density within each PSD. To determine this, we calculated the fractional distribution of PSD-95 protein within each PSD (see Methods for details), producing a map of molecular density (Fig. 3A). Protein concentration was inhomogeneous within individual PSDs, particularly in large PSDs (Fig. 3A). Using previous measurements of the number of PSD-95 molecules present at synapses (8, 12) (see SI Text), we converted our imaging data into an estimate of the absolute density of PSD-95 molecules in the PSD. The results of this analysis predict a mean density of 4.3 ± 0.4 PSD-95 molecules per 1000 nm2 in the PSD, or a center-to-center spacing of 17 ± 1 nm, remarkably consistent with the 10 nm size of purified PSD-95 (22) and the nearest-neighbor spacing of 13 nm measured by electron tomography (17), and supporting the notion of a dense arrangement of PSD-95 molecules capable of binding one another either directly or through minimal intermediates. Notably, the pattern of molecular density within PSDs changed over time, with localized concentration and dispersion of PSD-95 occurring within PSD subdomains over periods of minutes (Fig. 3B). PSD intermolecular positioning is thus inhomogeneous and malleable, and morphological plasticity of established PSDs involves reorganization of internal structure.

Fig. 3.
Actin drives spatial fluctuations of PSD-95 molecular density in individual PSDs. (A) Fractional protein density maps of single PSDs from several cells. The color scale shows the proportion of total molecules per 1,000 nm2. (Spatial scale bar: 1 μm.) ...

We next determined whether the actin cytoskeleton regulates PSD structural plasticity. Depolymerization of actin with latrunculin A abruptly halted ongoing PSD morphological changes (Fig. 3C and Movie S6). Cessation of PSD dynamics was not accompanied by a net change in EF (Fig. 3D), indicating that stabilization was not a relaxation toward a low energy or default state. Rather, a rapid drop in the CV of EF for individual PSDs (Fig. 3E) and a strongly reduced fraction of PSDs occupying dynamic Mode 1 (Fig. 3F) indicated that each PSD was quickly frozen in place upon actin depolymerization. As a result, the ongoing fluctuations of protein density within each PSD were reduced (Fig. 3G). Stabilization of the actin cytoskeleton with jasplakinolide also promptly stabilized PSD structure (Fig. S2 and Movie S7). Although actin regulates spine shape and motility, we detected no correlation between spine dynamics and PSD restructuring (Fig. S3, Additional Discussion in SI Text). We conclude that actin remodeling is required for ongoing PSD structural alteration.

Actin-dependent restructuring of the PSD could arise from the removal and addition of molecules at spatially distinct points (Fig. S4A). However, we found that the exchange of PSD-95-GFP was slow at the PSD (Fig. S4 B–D), consistent with previous work (22, 25, 26, 31). Preincubation of neurons with jasplakinolide or latrunculin for 15 min did not affect the low rate or extent of PSD-95-GFP fluorescence recovery (Fig. S4 E–G), demonstrating that, unlike PSD morphological dynamics and protein redistribution (Fig. 3 C–G), molecular exchange of this core PSD protein is actin-independent (25). The different rates and actin dependence of PSD restructuring compared to PSD-95 molecular exchange suggest that spatially distinct zones of molecular addition and removal do not account for PSD shape change.

Limited Mobility of PSD-95 While Bound in the PSD.

The above experiments establish that molecules are long-lived within the PSD, the shape of which is plastic. There are 2 general models for the architecture of such a structure (Fig. 4A): a bordered assembly with free internal movement (i.e., a fence or corral) or a flexible structure with fixed molecular positions (i.e., a grid or matrix). We used the formalism developed by Carrero et al. (32) to predict how quickly, within μm-sized structures such as the PSD, molecules of different mobility will spatially equilibrate. By solving the model using values for the expected diffusion coefficient D of different types of proteins, we found that freely diffusing, cytosolic molecules redistribute over the course of only a few tens of milliseconds (Fig. S5). Proteins such as PSD-95 that are attached to the membrane by palmitoylation or other lipid modifications are predicted to reach equilibrium distribution within 1 sec, and transmembrane proteins are expected to do so within seconds (32, 33) (Fig. S5). Because little information exists regarding molecular motion internal to the PSD, we developed a set of optical approaches to examine mobility of PSD-95 within individual PSDs spanning time scales of milliseconds to minutes.

Fig. 4.
Limited mixing of molecules within the PSD. (A) Two alternative models of PSD structure: a “corral” that contains movable molecules within a defined border (Left), or a “matrix” of molecules held in fixed positions within ...

To assay protein movement within the PSD at subsecond resolution, we measured FRAP by scanning along a single line through single PSDs at 200 Hz for 20 sec. Scanning laser power was low so that fluorescence intensity of unbleached PSDs remained stable (Fig. 4B). When the laser intensity was raised briefly along the entire extent of the PSD, the whole structure was bleached and little or no recovery was observed, consistent with the slow recovery shown in Fig. S4. To examine mobility within the complex, photobleaching was restricted to 1 side of the PSD (Fig. 4B). Surprisingly, the recovery of fluorescence in the bleached subregion was minimal over the extent of the experiment (Fig. 4B). XY scans before and after the line-scan confirmed that photobleaching was restricted to 1 side of the PSD, and directly demonstrated a large remaining pool of fluorescent PSD-95-GFP which did not equilibrate with the bleached molecules (Fig. 4C). The distribution of fluorescence intensity across the PSD remained essentially constant after bleaching (Fig. 4D). On average, recovery of fluorescence proceeded neither faster nor to a greater extent in the bleached fraction of partially bleached PSDs than in fully bleached PSDs (Fig. 4E; extent of recovery 8 ± 2% full vs. 10 ± 3% partial). In contrast, when PSD-95-GFP was expressed in glia, where it is palmitoylated and targeted to the plasma membrane (34), photobleaching with the same protocol revealed a rapid biphasic recovery with a half-recovery time of 1.7 sec (Fig. 4E; extent of recovery 77 ± 8%), consistent with our modeling results and D ≈ 0.1 μm2/sec (Fig. S5).

To assess movement within PSDs on time scales longer than a few seconds, we developed a fluorescence overlay photobleaching protocol. Coexpression of both cerulean- and citrine-tagged versions of PSD-95 allowed us to define the contour of the PSD with PSD-95-cerulean while photobleaching PSD-95-citrine in a visualized subregion of the PSD (Fig. 4F). After subregion photobleaching, the unequal distribution of fluorescence remained over several minutes, even in PSDs undergoing distinct changes in morphology (Fig. 4F and Fig. S6). The profile of unbleached citrine fluorescence along the length of the PSD changed little over this time (Fig. 4G).

To confirm the near-immobility of PSD-95 without relying on photobleaching, we co-expressed versions of PSD-95 tagged with mCherry and with photoactivatable GFP (PAGFP), and targeted portions of visualized PSDs for PAGFP photoactivation using a pulsed IR laser (Fig. 4H). For at least 60 sec following such microphotoactivation, photoactivated PSD-95-PAGFP molecules did not diffuse through the PSD, the overall shape of which could be measured following global activation (Fig. 4 H and I and Fig. S7). In a few cases, we were able to carry these approaches to much longer times, and found that the lack of mobility of PSD-95-citrine protein within the PSD persisted for as long as 30 min (Fig. S6E). By this point, recovery of fluorescence within the bleached region of the PSD appeared no greater than expected due to overall exchange rates in and out of the PSD. In total, this suite of results indicates that resident scaffold proteins exhibit little movement or mixing within the PSD, and show directly that molecularly distinct subdomains of the postsynaptic specialization can persist for long periods.

Discussion

We have analyzed two novel characteristics of PSD internal behavior. First, despite their long-term stability, individual PSDs have a continuously dynamic structure. This result suggests that the PSD is not a rigid scaffold or platform, because the structure is inherently flexible. This plasticity of PSD morphology and the internal distribution of PSD-95 molecules is driven by the actin cytoskeleton over the time scale of seconds to minutes, but not by molecular exchange, which is much slower and actin-independent. Second, high-resolution optical tagging of PSD subregions revealed very little scaffold protein movement within the PSD itself, providing direct insight to the internal architecture of this prototypical multiprotein ensemble. One model which reconciles these seemingly contradictory aspects of PSD architecture considers the PSD as a flexible yet topologically stable matrix (Fig. 5). This model reflects that we did not detect molecular mixing in our experiments. However, it remains possible that strong actin-driven deformation of PSDs (e.g., during the dynamic Mode 1 defined in Fig. 2) eventually drives breakdown of the matrix and mixing of PSD constituents over longer time scales than we have examined. It will be important for future studies to examine whether different forms of activity-regulated synaptic plasticity require a large-scale reorganization of the PSD network, and whether PSD integrity persists by addition or deletion of matrix elements at internally stable coordinates.

Fig. 5.
Flexible matrix model of PSD architecture. (Left) Scaffold proteins (yellow) establish matrix coordinates within the larger PSD complex (gray). (Right) Components of the matrix can be added (green) or removed (red), and this molecular exchange takes place ...

The Organization of Information Storage Within the Synapse.

Here we have shown that the core PSD scaffold protein PSD-95 forms an enduring, spatially stable matrix within the PSD with extremely limited mixing of molecules over submicron distances. A corollary of these findings is that exchange, addition, or removal of PSD-95 scaffold elements can occur at independent matrix positions, thereby establishing a Cartesian coordinate system, or molecular map, for organizing synaptic nanoarchitecture. Due to the limits of confocal microscopy, we cannot fully resolve the scale of the synaptic spatial coordinate system, and the nodes of this matrix may comprise either single molecules or larger protein modules. It is tempting to speculate that the nodes of the PSD matrix correspond to the ≤100 nm subsynaptic domains in which AMPA receptors are confined (35). Emerging superresolution microscopy techniques (36, 37) may in the future be able to provide live-cell views of PSD organization at sufficiently high resolution to distinguish these nanometer-scale complexes.

The regulated positioning and estimated molecular spacing and density of PSD-95 within the PSD reported here are consistent with a direct role for this protein and its family members in establishing the synaptic matrix (38). Indeed, recent EM tomography (17) has confirmed the distribution of PSD-95 spaced throughout the PSD, and suggested that intermolecular spacing of PSD-95 may be established through intermediate scaffolds (17). Such intermediates provide an attractive mechanism to flexibly link nodes of the matrix in Fig. 5. It will be important to assess whether these or other oligomeric interactions of PSD proteins (22, 39, 40) confer stability to the complex as a whole or to subsets of proteins.

A caveat of many live-cell approaches is the reliance on exogenous PSD-95, which is known to enlarge and strengthen synapses, occlude LTP, and enhance LTD (41). However, dynamics of PSD morphology were not sensitive to PSD-95-GFP expression level (data not shown) and were quantitatively preserved when endogenous PSD-95 was knocked down via shRNA and rescued with protein bearing silent mutations (Fig. S8). Further experiments using a variety of PSD proteins will help quantify morphodynamic characteristics in synapses of differing strength, developmental status, and history of plasticity.

Morphing of the Synapse Instead of the Spine.

Although the PSD comprises only a small part of a dendritic spine, changes in spine shape have been increasingly used as a readout for synaptic plasticity. However, inferring synapse functional state from spine geometry is at best indirect and may even at times be misleading (42, 43; also see SI Text, Additional Discussion). In this work, we have taken a new approach to obtain a more direct observation of synaptic dynamics. We reasoned that changes to the PSD itself are highly likely to reflect altered synapse function. PSD-95 is tightly associated with the membrane by N-terminal palmitoylation and by PDZ interactions with NMDA receptors, transmembrane AMPA receptor regulatory proteins (TARPs), and synaptic adhesion molecules (17, 44, 45). The deformation of the PSD that we have documented is thus likely accompanied by localized distortion of the postsynaptic membrane, the shape of the synaptic cleft, or the alignment of the presynaptic and postsynaptic membranes. Indeed, the strong enrichment and high turnover rates of actin and cytoskeletal proteins both in the terminal and in the spine (46) suggest that synapses are subject to substantial, continuous mechanical forces. Gephyrin scaffolds at inhibitory synapses are similarly subject to continuous perturbation by both actin and microtubules (47). Although the scarcity of microtubules in spines suggests that they have little direct influence on PSD morphology, our results make it clear that actin polymerization controls PSD morphological change. One possible mechanism of this control is that direct cytoskeletal connections to the PSD (48) localize spine actin polymerization and provide the propulsive force deforming the PSD.

Synaptic Function Controlled Without Altering Synaptic Protein Content.

A notable aspect of the flexible matrix architecture of the synapse defined here is that morphological elasticity from stretching or compressing the overall scaffold provides a possible means to control local protein density independent of protein exchange or addition. Such “breathing” of a scaffold matrix could serve to dynamically adjust the alignment of bound postsynaptic receptors with sites of presynaptic vesicle fusion. This alignment may be functionally significant, as numerical modeling suggests that glutamate concentration decays rapidly away from the site of vesicle fusion, meaning that small shifts in AMPA receptor position can create large changes in the amplitude and kinetics of postsynaptic responses (2, 50). Similarly, actin-driven shifts in scaffold positioning may regulate the local density of signaling molecules bound to PSD scaffolds, potentially activating or terminating signaling within PSD subdomains. The flexible matrix model described provides a rational basis for future investigation into the nanomolecular organization and dynamics of glutamatergic synapses.

Methods and SI Text.

Neurons were cultured from E18 rat hippocampus as described in ref. 51. Except where mentioned, cells after 3 to 6 weeks in culture were transfected with Lipofectamine 2000 (Invitrogen) (51). PSD morphology was observed using a Yokogawa CSU spinning disk confocal (51), and 3D stacks were projected to 2D before analysis principally in Metamorph. Linescan FRAP, whole-spine photoactivation, and overlay photobleaching were carried out on a Leica SP2 AOBS. Two-photon photoactivation of PSD subregions was performed on a Zeiss 510 Meta NLO equipped with a Coherent MaiTai Ti:Sapphire laser tuned to 820 nm.

Complete methods, referenced movies, supplemental figures, and additional discussion can be found in the SI Text, SI Methods, Figs. S1–S8, and Movies S1–S7.

Supplementary Material

Supporting Information:

Acknowledgments.

This work was supported by National Institutes of Health grants R01 AG024492, R01 NS047574, and NS039402 to M.D.E, and NIMH R01 MH080046 and a NARSAD Young Investigator Award to T.A.B. M.D.E. is an Investigator of the Howard Hughes Medical Institute. We thank I. Lebedeva, H. Zhang, and H. Kong for expert technical assistance, and M. Bond, K. Condon, N. Frost, T. Helton, J. Hernandez, M. Kennedy, M. Pucak, S. Raghavachari, and S. Thompson for helpful discussions and comments on the manuscript. We thank D. Bredt for the gifts of PSD-95-GFP and stargazin-GFP, E. Kim for HA-shank, R.Y. Tsien for mRFP, citrine, and mCherry, J. Lippincott-Schwartz for PA-GFP, M. Rizzo and D. Piston for cerulean, and J. Bear for pLL3.7.

Footnotes

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

This article contains supporting information online at www.pnas.org/cgi/content/full/0711669105/DCSupplemental.

References

1. Choquet D, Triller A. The role of receptor diffusion in the organization of the postsynaptic membrane. Nat Rev Neurosci. 2003;4:251–265. [PubMed]
2. Lisman J, Raghavachari S. A unified model of the presynaptic and postsynaptic changes during LTP at CA1 synapses. Sci STKE. 2006;356:re11. [PubMed]
3. Shepherd JD, Huganir RL. The cell biology of synaptic plasticity: AMPA receptor trafficking. Annu Rev Cell Dev Biol. 2007;23:613–643. [PubMed]
4. Murthy VN, Schikorski T, Stevens CF, Zhu Y. Inactivity produces increases in neurotransmitter release and synapse size. Neuron. 2001;32:673–682. [PubMed]
5. Harris KM, Stevens JK. Dendritic spines of CA 1 pyramidal cells in the rat hippocampus: Serial electron microscopy with reference to their biophysical characteristics. J Neurosci. 1989;9:2982–2997. [PubMed]
6. Takumi Y, Ramirez-Leon V, Laake P, Rinvik E, Ottersen OP. Different modes of expression of AMPA and NMDA receptors in hippocampal synapses. Nat Neurosci. 1999;2:618–624. [PubMed]
7. Kharazia VN, Weinberg RJ. ImmunoGold localization of AMPA and NMDA receptors in somatic sensory cortex of albino rat. J Comp Neurol. 1999;412:292–302. [PubMed]
8. Sugiyama Y, Kawabata I, Sobue K, Okabe S. Determination of absolute protein numbers in single synapses by a GFP-based calibration technique. Nat Methods. 2005;2:677–684. [PubMed]
9. Scheff SW, Price DA. Synaptic pathology in Alzheimer's disease: A review of ultrastructural studies. Neurobiol Aging. 2003;24:1029–1046. [PubMed]
10. Inoue A, Okabe S. The dynamic organization of postsynaptic proteins: Translocating molecules regulate synaptic function. Curr Opin Neurobiol. 2003;13:332–340. [PubMed]
11. Nicholson DA, Yoshida R, Berry RW, Gallagher M, Geinisman Y. Reduction in size of perforated postsynaptic densities in hippocampal axospinous synapses and age-related spatial learning impairments. J Neurosci. 2004;24:7648–7653. [PubMed]
12. Chen X, et al. Mass of the postsynaptic density and enumeration of three key molecules. Proc Natl Acad Sci USA. 2005;102:11551–11556. [PMC free article] [PubMed]
13. Ehlers MD. Activity level controls postsynaptic composition and signaling via the ubiquitin-proteasome system. Nat Neurosci. 2003;6:231–242. [PubMed]
14. Husi H, Ward MA, Choudhary JS, Blackstock WP, Grant SG. Proteomic analysis of NMDA receptor-adhesion protein signaling complexes. Nat Neurosci. 2000;3:661–669. [PubMed]
15. Sheng M, Hoogenraad CC. The postsynaptic architecture of excitatory synapses: A more quantitative view. Annu Rev Biochem. 2007;76:823–847. [PubMed]
16. Petersen JD, et al. Distribution of postsynaptic density (PSD)-95 and Ca2+/calmodulin-dependent protein kinase II at the PSD. J Neurosci. 2003;23:11270–11278. [PubMed]
17. Chen X, et al. Organization of the core structure of the postsynaptic density. Proc Natl Acad Sci USA. 2008;105:4453–4458. [PMC free article] [PubMed]
18. Valtschanoff JG, Weinberg RJ. Laminar organization of the NMDA receptor complex within the postsynaptic density. J Neurosci. 2001;21:1211–1217. [PubMed]
19. Bresler T, et al. Postsynaptic density assembly is fundamentally different from presynaptic active zone assembly. J Neurosci. 2004;24:1507–1520. [PubMed]
20. Ebihara T, Kawabata I, Usui S, Sobue K, Okabe S. Synchronized formation and remodeling of postsynaptic densities: Long-term visualization of hippocampal neurons expressing postsynaptic density proteins tagged with green fluorescent protein. J Neurosci. 2003;23:2170–2181. [PubMed]
21. Marrs GS, Green SH, Dailey ME. Rapid formation and remodeling of postsynaptic densities in developing dendrites. Nat Neurosci. 2001;4:1006–1013. [PubMed]
22. Nakagawa T, et al. Quaternary structure, protein dynamics, and synaptic function of SAP97 controlled by L27 domain interactions. Neuron. 2004;44:453–467. [PubMed]
23. Gerrow K, et al. A preformed complex of postsynaptic proteins is involved in excitatory synapse development. Neuron. 2006;49:547–562. [PubMed]
24. Gray NW, Weimer RM, Bureau I, Svoboda K. Rapid redistribution of synaptic PSD-95 in the neocortex in vivo. PLoS Biol. 2006;4:2065–2075. [PMC free article] [PubMed]
25. Kuriu T, Inoue A, Bito H, Sobue K, Okabe S. Differential control of postsynaptic density scaffolds via actin-dependent and -independent mechanisms. J Neurosci. 2006;26:7693–7706. [PubMed]
26. Sharma K, Fong DK, Craig AM. Postsynaptic protein mobility in dendritic spines: Long-term regulation by synaptic NMDA receptor activation. Mol Cell Neurosci. 2006;31:702–712. [PubMed]
27. Turrigiano GG, Nelson SB. Hebb and homeostasis in neuronal plasticity. Curr Opin Neurobiol. 2000;10:358–364. [PubMed]
28. Kim MJ, et al. Synaptic accumulation of PSD-95 and synaptic function regulated by phosphorylation of serine-295 of PSD-95. Neuron. 2007;56:488–502. [PubMed]
29. Smith KE, Gibson ES, Dell'Acqua ML. cAMP-dependent protein kinase postsynaptic localization regulated by NMDA receptor activation through translocation of an A-kinase anchoring protein scaffold protein. J Neurosci. 2006;26:2391–2402. [PubMed]
30. Tsuriel S, et al. Local sharing as a predominant determinant of synaptic matrix molecular dynamics. PLoS Biol. 2006;4:1572–1587. [PMC free article] [PubMed]
31. Okabe S, Urushido T, Konno D, Okado H, Sobue K. Rapid redistribution of the postsynaptic density protein PSD-Zip45 (Homer 1c) and its differential regulation by NMDA receptors and calcium channels. J Neurosci. 2001;21:9561–9571. [PubMed]
32. Carrero G, McDonald D, Crawford E, de Vries G, Hendzel MJ. Using FRAP and mathematical modeling to determine the in vivo kinetics of nuclear proteins. Methods. 2003;29:14–28. [PubMed]
33. Sprague BL, Pego RL, Stavreva DA, McNally JG. Analysis of binding reactions by fluorescence recovery after photobleaching. Biophys J. 2004;86:3473–3495. [PMC free article] [PubMed]
34. Topinka JR, Bredt DS. N-terminal palmitoylation of PSD-95 regulates association with cell membranes and interaction with K+ channel Kv1.4. Neuron. 1998;20:125–34. [PubMed]
35. Ehlers MD, Heine M, Groc L, Lee MC, Choquet D. Diffusional trapping of GluR1 AMPA receptors by input-specific synaptic activity. Neuron. 2007;54:447–460. [PMC free article] [PubMed]
36. Betzig E, et al. Imaging intracellular fluorescent proteins at nanometer resolution. Science. 2006;313:1642–1645. [PubMed]
37. Sieber JJ, et al. Anatomy and dynamics of a supramolecular membrane protein cluster. Science. 2007;317:1072–1076. [PubMed]
38. Ehrlich I, Klein M, Rumpel S, Malinow R. PSD-95 is required for activity-driven synapse stabilization. Proc Natl Acad Sci USA. 2007;104:4176–4181. [PMC free article] [PubMed]
39. McGee AW, et al. Structure of the SH3-guanylate kinase module from PSD-95 suggests a mechanism for regulated assembly of MAGUK scaffolding proteins. Mol Cell. 2001;8:1291–1301. [PubMed]
40. Baron MK, et al. An architectural framework that may lie at the core of the postsynaptic density. Science. 2006;311:531–535. [PubMed]
41. Beique JC, Andrade R. PSD-95 regulates synaptic transmission and plasticity in rat cerebral cortex. J Physiol. 2003;546:859–867. [PMC free article] [PubMed]
42. Sdrulla AD, Linden DJ. Double dissociation between long-term depression and dendritic spine morphology in cerebellar Purkinje cells. Nat Neurosci. 2007;10:546–548. [PubMed]
43. Wang XB, Yang Y, Zhou Q. Independent expression of synaptic and morphological plasticity associated with long-term depression. J Neurosci. 2007;27:12419–12429. [PubMed]
44. Chen L, et al. Stargazin regulates synaptic targeting of AMPA receptors by two distinct mechanisms. Nature. 2000;408:936–943. [PubMed]
45. el-Husseini Ael D, Bredt DS. Protein palmitoylation: A regulator of neuronal development and function. Nat Rev Neurosci. 2002;3:791–802. [PubMed]
46. Star EN, Kwiatkowski DJ, Murthy VN. Rapid turnover of actin in dendritic spines and its regulation by activity. Nat Neurosci. 2002;5:239–246. [PubMed]
47. Hanus C, Ehrensperger M-V, Triller A. Activity-dependent movements of postsynaptic scaffolds at inhibitory synapses. J Neurosci. 2006;26:4586–4595. [PubMed]
48. Ethell IM, Pasquale EB. Molecular mechanisms of dendritic spine development and remodeling. Prog Neurobiol. 2005;75:161–205. [PubMed]
49. Matus A. Actin-based plasticity in dendritic spines. Science. 2000;290:754–758. [PubMed]
50. Raghavachari S, Lisman JE. Properties of quantal transmission at CA1 synapses. J Neurophysiol. 2004;92:2456–2467. [PubMed]
51. Blanpied TA, Scott DB, Ehlers MD. Regulation and dynamics of clathrin at specialized endocytic zones in dendrites and spines. Neuron. 2002;36:435–449. [PubMed]

Articles from Proceedings of the National Academy of Sciences of the United States of America are provided here courtesy of National Academy of Sciences
PubReader format: click here to try

Formats:

Related citations in PubMed

See reviews...See all...

Links

Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...