NCBI Bookshelf. A service of the National Library of Medicine, National Institutes of Health.

Kittler JT, Moss SJ, editors. The Dynamic Synapse: Molecular Methods in Ionotropic Receptor Biology. Boca Raton (FL): CRC Press/Taylor & Francis; 2006.

Cover of The Dynamic Synapse

The Dynamic Synapse: Molecular Methods in Ionotropic Receptor Biology.

Show details

Chapter 9Receptor Dynamics at the Cell Surface Studied Using Functional Tagging

and .

9.1. INTRODUCTION

Heisenberg’s Uncertainty Principle, originating from quantum mechanics, can also be appropriately applied to the study of receptor dynamics in cell membranes. Essentially, the “uncertainty” is introduced because to measure or track receptor movements in cell membranes, we really need to tag the proteins, which in itself could alter receptor mobility. The nature of the tag might involve a sequence of amino acids inserted into the protein (structural modification), or simply the addition of an antibody to a native epitope on the receptor. In either example, the accuracy of measurements of mobility will innately have a degree of uncertainty associated with them. The key objective of any method employed to measure receptor movements is to minimize this uncertainty while maximizing the resolution fidelity for an appropriate population of receptors.

To date, the principal and most technically accessible methods for monitoring the dynamic movement of receptors within the membrane of neuronal or other cells most commonly involves some form of biochemical or optical procedure. Optical approaches have one critical advantage in that they provide reasonably high temporal resolution, especially when images of membrane receptors or clusters of receptors are obtained from live cells that are monitored using organic fluorophores or quantum dots, in concert with the rapid acquisition capabilities of confocal microscopy. However, both biochemical and optical techniques require certain assumptions to be made before they can impart information about the movements of functional receptors in the cell membrane. Such techniques will, to some extent, inevitably include receptors from several pools (e.g., cell surface membrane and intracellular pools), some of which might be nonfunctional, or have limited involvement in maintaining the appropriate excitable state of the neurone. The functional approaches described in this chapter use a different strategy based on electrophysiological methods. This provides a measure of the current which results only from the binding of activating ligands to their respective ligand-gated ion channels (LGIC) thereby resolving only those receptors that actually contribute to the excitable state of the neuron. This can include both synaptic and extrasynaptic pools of functional receptors but, importantly, it will discount submembranous receptors. As such, the influence these tagged receptors contribute to the metaplastic changes (potentiation or depression) of the synaptic response can, in principle, be readily monitored. As we will see, this method of functional tagging requires that particular receptor isoforms carry either a biophysical or pharmacological reporter, or tag, that can be readily studied in live cells, and that such reporters provide distinctive functional signatures. This could be manifest by the activation, modulation or ablation of receptor function, depending on the tag and its ligand as described in the proceeding sections.

9.2. PERSPECTIVE ON NONFUNCTIONAL RECEPTOR TAGGING TECHNIQUES

Why should we seek alternative receptor tracking strategies to the many optical, biochemical and other nonfunctional methods that follow the life-cycle of membrane receptors? The simplest answer to this question is that certain procedures are more suitable to address specific questions with regard to the trafficking of membrane proteins and, importantly, not all these methods selectively monitor the same parameters. In many cases, several of these techniques are complementary.

The application of a functional tag to a receptor (also referred to as “electrophysiological tagging”) permits experiments that monitor the stability, biophysical properties and pharmacological profile of a functionally competent population of receptors, which are resident in the cell membrane, behaving functionally as they would in vivo. The functional tag therefore excludes any direct contributions that might be made to the overall monitoring of receptor movement by immature, dysfunctional or submembranous receptors. Moreover, by applying tags to certain specific populations of receptors, or by enabling a tag to become visible only during particular functional states of the receptor, the relative trafficking of synaptic receptors, as distinct from extrasynaptic receptors, can be deduced. This is an important difference between functional tagging and nonfunctional biochemical or optical methods. The advent of organic dyes such as FM4-64 [1,2] and related products, which can be uptaken into synaptic vesicles, has attempted to rectify this to some extent by permitting only the identification of active synapses, although this fluorescent dye does not identify the movement of functionally active receptors [3]. Calcium imaging of dendritic spines offers another optical approach to the measure of functional receptor activity, although this can have spatio-temporal resolution limitations and still represents a second-order response following receptor activation. It has yet to be fully exploited to monitor receptor movements in and around the cell membrane.

As receptor tracking experiments are often required to monitor movements in the membrane over relatively short time periods, the temporal resolution of biochemical and fixed staining procedures can be a limitation. The practicalities of conducting pulse-chase experiments (either with antibodies conjugated to radioactive tracers, avidin or specific organic fluorophores) is generally not conducive to following the fast movements of membrane-delimited proteins, which occur minute-by-minute under conditions of both basal and stimulated neuronal activity. As far as immunocytochemistry is concerned, temporal issues have been resolved to some extent through novel protein modifications, such as the introduction of cleavable epitopes such as thrombin and the haemagglutinin tag [4]; fusions with GFP; pH-sensitive GFPs (pHlourins) [5,6]; cysteine-biarsenicals [7–9]; and the introduction of high-affinity, irreversible binding sites for high potency toxins, e.g., α-bungarotoxin [10]. Many of these epitopes or tags can be used in live staining procedures. Perhaps the most useful of these are the pHluorins, particularly ecliptic pHluorin, which uniquely loses fluorescence at one excitation wavelength when placed in an acidic pH environment that is normally only experienced by internalized receptors rather than their cell surface counterparts. Thus, this pHluorin should faithfully report the movements of principally cell surface receptors, although certain trans-Golgi compartments, from which synthesized receptors originate, have sufficiently neutral pH to permit fluorescence. Thus, with few exceptions, fluorescently labeled receptors that are internalized or are being trafficked to the cell surface are usually difficult to unequivocally distinguish from those that only reside at the cell surface.

Antibody labeling techniques have further limitations. Antibodies are many orders of magnitude larger than the receptor they are labeling. Generally, this disparity in size is exacerbated by the requirement for a secondary antibody to be linked to an organic fluorophore for visualization and, indeed, this disparity is exacerbated further with quantum dot coupling. In addition, the primary antibody needs to be raised to external receptor epitopes to distinguish only cell surface receptors and avoiding the need to membrane permeabilize to allow the antibody to gain access to intracellular epitopes, which would make intracellular and membrane receptor pools indistinguishable. Furthermore, dual labeling of receptors in concert with specific synaptic marker proteins only indicates the general “proximity,” not absolute apposition, of a receptor to a synaptic site. One further caveat also suggests that by no means are all synaptically labeled receptors functionally active [3]. Yet further ambiguity is raised because synaptic and extrasynaptic receptor classification at the optical level is usually ascribed on the basis of immunofluorescent puncta size, although what part individual receptor-antibody-fluorophore complexes represents in a single punctum and how this reflects on the potential number of receptors per punctum is unknown. Further, under conditions of prolonged exposure to excitable wavelengths of laser light, such as occurs with repetitive line scanning of the same region of interest (a common technique used to optimize an image and to perform time-series studies), certain organic fluorophores are susceptible to extensive bleaching. As such, unintentional fluorescence loss in photobleaching (FLIP) or fluourescence recovery after photobleaching (FRAP) measurements can give the false impression of receptor mobility.

9.3. EARLY STUDIES USING FUNCTIONAL TAGGING

Relatively few examples are found of the use of functional tags to study the membrane movements of neurophysiologically important proteins. Because of technological limitations, many early examples were unable to introduce a tag into a precise part of the receptor structure. As such, early experiments, although often ingenious, were somewhat unrefined as, for example, in experiments designed to assess the mobility of voltage-gated potassium and sodium channels within the plasma membrane of frog muscle fibers. Here, a patch pipette was used to measure membrane currents and then also to shine UV light onto the membrane patch to cause photo destruction of the ion channels (and, presumably, other surface proteins also) [11]. By subsequently monitoring the faster recovery of membrane potassium currents relative to sodium currents, potassium channels were concluded to possess greater mobility than their sodium channel counterparts.

An earlier yet somewhat more sophisticated approach relied on the use of an inherent pharmacological reporter within the muscle nicotinic acetylcholine receptor (AChR), which is irreversibly inhibited by α-bungarotoxin with nanomolar affinity. The density of AChR on the myotomal muscle cell surface of Xenopus tadpoles was monitored by the regular iontophoretic application of ACh, which caused membrane depolarizations [12]. Local inactivation of these functional AChRs by α-bungarotoxin and monitoring the recovery of the depolarizations (Figure 9.1a and Figure 9.1b) permitted a determination of the diffusion coefficient (D) for these receptors. The value of D (1.5 × 109 to 4.0 × 109 cm2/sec) implied that simple diffusion-based redistribution is the likeliest mechanism for receptor movement during synaptogenesis.

FIGURE 9.1. Early form of receptor tagging.

FIGURE 9.1

Early form of receptor tagging. (a) Intracellular recording (V) from a Xenopus tadpole myotomal muscle cell showing repetitive localized application of acetylcholine (ACh) in the absence or presence of focally-applied α-bungarotoxin (BTX). (b) (more...)

An approximate estimate of how long (t) it takes a particle to diffuse a given distance (d) knowing its diffusion coefficient (D) can be found from: t1/2 – ≈ (d1/2)2/D. Thus, for a diffusion coefficient of 2.5 × 109 cm2/sec, to move a quarter of the way around a muscle fiber of diameter 30 μm will take approximately 37 min. For Young and Poo’s study [12], half recovery following α-bungarotoxin was reached for similar sized fibers in approximately 21 min. Therefore, diffusion alone was sufficient to explain the rate of recovery observed.

The technique successfully exploited another specific pharmacological reporter of the muscle AChR, concanavalin-A, which was used to immobilize AChRs and thus block diffusion (Figure 9.1c). This agent prevented the diffusion of toxin-blocked and unblocked receptors affecting the dynamics of recovery. The mechanism of this recovery represented the first electrophysiological demonstration that functional receptors were able to diffuse freely within the membrane. From these studies, the suggestion of a “diffusion trap” model for protein movement within the membrane originated. This entailed the region of innervation on a post-synaptic membrane serving as a sink, or trap, for functional receptors that are readily diffusing in the membrane, thus leading to their initial concentration during synaptogenesis and subsequent maintenance at mature synapses. Recently, this concept has again found favor in explaining the movements of neurotransmitter receptors in cell membranes [13].

9.4. THE NATURE OF THE EPITOPE: CRITERIA FOR SELECTING A FUNCTIONAL TAG

Trying and achieving several of the following criteria is desirable when selecting or adopting a functional tag to monitor the movement of surface receptors. The main objective is to monitor the receptors without unduly affecting their behavior in the membrane by the inclusion of a functional tag. Importantly, the epitope could be an innate part of the native receptor, which is functionally susceptible to the binding of a ligand, for example, or it could be part of the receptor that can be conveniently engineered by mutagenesis and reintroduced into a cell to become susceptible to another ligand that might not normally associate with that protein. Although either approach is useful, other criteria are worthy of consideration.

Tag size

This attribute should be kept to a small size; large tags, particularly those that increase the size of the receptor in terms of bulk or mass, are more likely to have an impact on receptor movement and speed of diffusion in the membrane.

Silent tag

The incorporation of a tag alone should not affect the mobility, distribution or the function of the receptor. Thus, the tag should act in a passive capacity. Binding a ligand to the tag might then specifically alter the function of the receptor, which can be used as a monitor of receptor movement. Essentially, the tag must therefore remain silent until it is activated or becomes bound to another moiety.

Irreversible tag activation

To accurately follow the movements of tagged receptor proteins, tag activation should be irreversible or, at best, only very slowly reversible. This criterion will ensure that a tagged receptor that becomes functionally altered following ligand binding can be tracked faithfully until it is endocytosed without any confounding observations resulting from the ligand dissociating from the tag and the receptor function altering as a result.

Receptor assembly

The inclusion of the tag must not interfere with subunit-subunit assembly for hetero- or homo-oligomeric receptors.

Innate tag

Where possible, to guarantee the characteristics listed above, it is better to use an innate tag, i.e., an epitope or binding site naturally present on the receptor. This use will ensure the least disruption to normal receptor function and trafficking. As is often the case, innate tags, apart from being sensitive to selective antisera, can be quite unsuitable for tracking receptor movement due to the inappropriateness of the binding ligand for such a function (i.e., the effect of the ligand might not be easy to monitor).

State-dependent tag

To gain maximum use of a functional tag, only revealing the epitope to a ligand during particular conformational states of the receptor is helpful. For example, if the tag is present in the ion channel, then only revealing this tag when the ion channel is activated is useful, thus rendering active channels susceptible to tagging by a suitable ligand but not their closed counterparts.

Rapid functional reporter

To provide immediate feedback on receptor movement, the binding of a ligand to an epitope and its subsequent effect on receptor function should be rapid or at least several orders of magnitude faster than the movement of receptors that are being tracked.

9.5. ADVANTAGES OF THE FUNCTIONAL TAG IN THE STUDY OF MOBILE RECEPTORS

The binding of a ligand to a receptor possessing an integral ion channel involves a characteristic signal transduction event involving the opening of the ion channel and flow of ions. This ionic current represents the normal physiological function of the receptor but, as such, it can also be used to act as a reporter for the receptor’s movement in real-time. The ability to measure and monitor currents through functional receptors is the principal property that makes the functional tagging approach a more physiological measurement than corresponding biochemical or optical techniques.

Generally, the phasic release of neurotransmitter from pre-synaptic specializations (in mature primary cultures, acute tissue slices, organotypic cultures or even primary and secondary cell line co-cultures) activates only post-synaptic receptors that are located directly apposed to the transmitter release sites. These phasic synaptic currents are transient in nature and their profile depends on the receptor subtype (subunit composition) present at synapses, their biophysical characteristics (i.e., activation, deactivation and desensitization rates, degree of rectification and so on), their state of modulation (e.g., phosphorylation) and the speed of removal of the transmitter by transporters or inactivating enzymes. These post-synaptic receptors can often be distinct in terms of composition and pharmacology from their perisynaptic and extrasynaptic equivalents that populate the remainder of the neuronal membrane. Extrasynaptic receptors that are persistently activated by neurotransmitter spillover from synapses or basal levels of transmitter from other sources (e.g., astrocytes) are responsible for tonic currents, which partly set the threshold at which action potentials are generated, thereby contributing to “signal integration” in the neuron [14–16].

For a particular receptor family or a receptor subtype within a single family, the presence of a functional tag allows the current flow through specified receptors to be manipulated using specific pharmacological agents, often inhibitors. For the agent to be maximally useful, its binding site can only be exposed once the channel is activated, thus making the use-dependent channel blocking agent a popular tool for functional tagging studies. The ideal, irreversible (covalent) binding reaction of a ligand to the epitope ensures that any recovery of function can only be attributable to the introduction (by recycling, exocytosis or lateral translocation) of unblocked receptors into the cell membrane.

The advantage of this type of block of receptors in a neuron is that the spontaneous and miniature synaptic currents, due to neurotransmitter release only at the synapse, can be studied in isolation from the extrasynaptic receptor pool, i.e., only the activated receptors at the synapse are susceptible to block. Once the functionally irreversible current “knockdown” is complete, any recovery of synaptic activity will be the result of unblocked receptors moving in from either extrasynaptic sites or from exocytotic pathways to displace these blocked synaptic receptors. By the nature of the available ligands, some receptors possess a native, inherent functional tag, e.g., α-amino-3-hydroxy-5-methyl-4-isoxazole propionic acid (AMPA) receptors and N-methyl-D-aspartate (NMDA) receptors, but others require to have such tags engineered into the protein, as is the case for the GABAA receptors (see section 9.8).

9.6. AMPA RECEPTOR RECTIFICATION AND SYNAPTIC “AMPAFICATION”

Early measurements of protein mobilities, revealed that integral membrane proteins such as rhodopsin have restricted movement in the plane of the cell membrane compared to being able to diffuse freely in the lipid bilayer [17]. Furthermore, measurements of AChR mobility at the neuromuscular junction revealed such a slow diffusion coefficient (D < 1014 cm2/sec) that monitoring movement was difficult with any accuracy [18,19].

Indeed, given such a small diffusion coefficient, the calculation in section 9.3 regarding the movement of AChRs around one-quarter of a muscle fiber would now take approximately 17.6 years. Thus, such receptors could be considered as static entities.

Now well-established is the fact that this stability of the receptors in the post-synaptic membrane is essential for effective intercellular communication, as the entrapment of mobile receptors permits the apposition of pre- and post-synaptic structures at neurotransmitter release sites. The slowness of some receptors to diffuse in the membrane has been unequivocally established as due to specific interactions between anchoring proteins and post-synaptic receptors in the post-synaptic density (PSD) [20–22].

Although anchored at the synapse, these receptors are able to exhibit mobility as observed during activity-dependent changes to the efficacy of synaptic transmission that underlie plasticity changes such as long-term potentiation (LTP) and long-term depression (LTD). The molecular basis for these changes could stem from changes to the channel conductance [23], alterations to the phosphorylation state of the receptor [24–27], as well as increased delivery of receptors to the synapse [28]. The first studies to correlate the effects of neuronal activity on the trafficking of AMPA receptors at individual synapses coupled observations of cell surface epitope tagged AMPA receptors and electrophysiological recordings of mEPSC events [29–31]. Subsequent functional tagging studies [32] clarified evidence from optical and biochemical data that the pool of membrane receptor proteins, rather than being rigidly corralled and retained as part of the PSD, is very dynamic, trafficking and targeting specific receptors into established synapses. As a consequence of this synaptic tuning, detectable changes to the functional signature of synaptic receptors could be recorded in response to stimulation.

Recording from cerebellar stellate cells, Lui and Cull-Candy [32] isolated AMPA-mediated excitatory post-synaptic currents (EPSCs) by minimally evoking the parallel fiber input in the presence of appropriate receptor blockers. The functional tag in this case exploited the susceptibility of AMPA receptor-derived EPSCs (which, importantly, lack the GluR2 subunit) to voltage-dependent block by the intracellular application of the polyamine, spermine. The net result is that while non-GluR2 subunit-containing AMPA receptors exhibit inward rectification due to their calcium permeability, inclusion of GluR2 confers relative calcium impermeability and no inward rectification [33–36]. This characteristic derives from the exclusive editing of the GluR2 subunit in the pore-lining region involving a Q/R mutation [37]. Interestingly, the GluR2 subunit is also the key to the susceptibility of AMPA receptors to other pharmacological agents. For instance, Joro spider toxin, pentobarbital and thiocyanate ions all differentially block AMPA-derived EPSCs depending on the presence of GluR2.

By measuring the degree of rectification for evoked EPSCs in the presence of spermine, any changes taking place to the functional properties of synaptic glutamate receptors in response to high-frequency stimulation could be monitored [32]. Synaptic currents, measured between 15 to 30 min after cessation of the stimulation, were seen to change from being predominantly inwardly-rectifying to linear (Figure 9.2). These results reflected a change in the AMPA receptor phenotype at the synapse due to an increased presence of the GluR2 subunit and emphasized the highly dynamic nature of the receptors in the PSD. However, the cellular origin of this new phenotype of synaptic receptor was not investigated but might have been as a result of increased protein expression, exocytotic delivery into the membrane from intra-cellular pools or in the membrane lateral translocation from extrasynaptic sources. The time frame for these changes suggested limited involvement of de novo protein expression. Thus, high-frequency stimulation can trigger the appearance of calcium-impermeable AMPA receptors at cerebellar stellate cell synapses that can be monitored as a functional change. Importantly, the same effect was not observed for low-frequency stimulation but could be emulated by bath application of glutamate or kainate. Thus, this is the first significant study to exploit an innate electrophysiological tag (i.e., the edited GluR2 subunit) to monitor the dynamics of a functional receptor. As an aside, this study also highlighted the importance of calcium entry through the AMPA receptor in inducing this subunit composition change, enabling a form of plasticity that is self-regulating because the expressed replacing synaptic receptors have much lower permeability to calcium.

FIGURE 9.2. Mobile AMPA receptors at glutamatergic synapses.

FIGURE 9.2

Mobile AMPA receptors at glutamatergic synapses. (a) After high-frequency stimulation (300 stimuli at 50 Hz), cerebellar stellate-cell EPSC amplitudes were reduced at –60 mV but increased at +40 mV holding potentials. (b) The inwardly-rectifying (more...)

Mediation of fast excitatory synaptic transmission in all parts of the central nervous system (CNS) is undertaken by AMPA receptors, whereas NMDA receptors are more crucial to a diverse range of developmental, physiological and pathological processes. Despite such disparate neurophysiological roles, these two receptor families are mutually dependent. This relationship is demonstrated early in development by the awakening of “silent” glutamatergic synapses whereby neuronal stimulation promotes the delivery of AMPA receptors (probably from extrasynaptic compartments, as well as intracellular pools) into the quiescent NMDA R-containing synapse causing their functional “awakening.” Even at later developmental stages, immunogold labeling techniques reveal that although NMDA receptors can be found in most mature synapses of the CA1 hippocampus, the same cannot be said of AMPA receptors [28,38,39]. However, changes in the plasticity of such mature pathways induced by stimulation are also underpinned by introducing functional AMPA receptors at synapses (AMPAfication) through the activation of NMDA receptors [40]. Thus, several parallels between synaptic maturation and LTP exist, principally involving the activity-driven delivery of AMPA receptors following NMDA receptor activation.

The dynamics of “silent” synapse activation and LTP have been addressed in a complementary study to that of Lui and Cull-Candy [32] using the same rectification property of GluR2-deficient receptors as a functional reporter. Hayashi et al. [41] over-expressed the GFP-tagged GluR1 AMPA receptor subunit in hippocampal slice neurons. Importantly, this caused the accumulation of largely homomeric GluR1 receptors in extrasynaptic regions of the dendrite, thus maintaining the linear rectification properties of endogenous GluR2-containing synaptic responses. Activation of CaMKII (which would occur following NMDA receptor activation and associated Ca2+ influx) caused the insertion of homomeric GluR1 into synapses, as detected by inward rectification of the EPSCs. Notably, pairwise recordings from GluR1-infected or noninfected cells in the absence of activated CaMKII failed to generate this switch. Further, an LTP induction protocol caused similar changes to the rectification of the EPSCs due to the import of GluR1 homomeric receptors into the synapse, which occurred 30 min after a stable potentiation period. Thus, phosphorylation events mediated by CaMKII appear to regulate the delivery of different phenotypes of synaptic AMPA receptors from extrasynaptic compartments into the synapse, changing the functional character of the synaptic response.

9.7. MK801 CHANNEL BLOCK EXPOSES THE TRANSIENT NATURE OF SYNAPTIC NMDA RECEPTORS

The NMDA receptor, unlike their AMPA-sensitive counterparts, were until recently considered to be relatively immobile elements of the PSD, being tightly anchored in the post-synaptic membrane [42]. Their resistance to detergent extraction from PSDs is testament to this [43,44], although, surprisingly, the half-life of NMDA receptors in cultured cerebellar granule cells is approximately only 1 day and not too dissimilar to that for AMPA receptors [45]. NMDA receptors are present at the synapse early in development and, until the activity-dependent importation of AMPA receptors into the synapse, constitute the “silent” synapse. The signal transduction cascades, mediated by Ca2+ influx through the NMDA receptor, are responsible not only for synapse formation but also for their modification and elimination [46,47]. Much of the plasticity of glutamatergic synapses has been attributed to the profound mobility of the AMPA receptor family but more recent evidence has underscored the contribution made by NMDA receptor turnover and trafficking [48–50].

The NMDA receptor open channel blocker, MK801, has proved a useful tool in the limited number of physiological studies of NMDA receptor dynamics in neurones. Whole-cell responses to NMDA are completely blocked by MK801 and show little recovery thereafter [51], suggesting limited replenishment of surface receptors. Interestingly, exposure of neurones to a constitutively active form of protein kinase C increased the exocytosis of unblocked, functional receptors to the membrane by nearly four-fold, thus highlighting the ability of these receptors to respond to stimuli and readily modify their numbers at the cell surface. A subsequent exploitation of MK801 was undertaken by Tovar and Westbrook [52] who, for the first time in a physiological study, were able to suggest that NMDA receptors move quite freely within the plane of the lipid bilayer between synaptic and extrasynaptic sites.

Using cultured hippocampal neurones forming autapses, all synaptic and extra-synaptic NMDA receptors were completely blocked by co-applying NMDA and MK801 to the whole cell. No recovery of NMDA receptor function was evident 30 min after MK801 washout, suggesting a very slow basal exocytosis rate for NMDA receptors (Figure 9.3a). However, when only synaptic receptors (measured as stimulated EPSCs) were blocked by MK801 (applied during the evoked EPSCs), a maximal recovery of about 40% in the synaptic response was seen within 20 min (Figure 9.3b). Because recovery from MK801 blockage was less than complete, only a limited reserve of NMDA receptors seems to be quickly mobilized into the synapse. This recovery of synaptic receptors could be accounted for by a number of scenarios, including dissociation of MK801 or an increase in the number of synaptic NMDA receptors. The latter could occur either at existing synapses, following new synapse formation or by an increase in the size of existing synapses. MK801 unbinding was discounted because exposing MK801 blocked synaptic receptors to AP5 (to prevent channel opening and MK801 unbinding) during washout saw the same level of current recovery. Moreover, insertion of new receptors was clearly not the route for recovery since the whole-cell responses did not recover. Latrunculin, which arrests dendritic spine mobility, also discounted active zone or pre-synaptic terminal migration as another possibility. Thus, the strongest candidate mechanism for synaptic receptor recovery was the lateral movement of unblocked extrasynaptic receptors within the membrane. Further experiments undertaken with another NMDA channel blocker, ketamine, revealed the true dynamic nature of these receptors, in that about 25% of the receptors on the entire cell surface entered a synapse within 5 min. This reinforces the diffusional rate constants seen for other ligand-gated ion channels in neuronal membranes [13,53–55] and also the high density of active synapses present on in vitro neurons, which could only be a fraction of the in vivo scenario. A further observation from these studies was that the number NMDA receptors moving into and out of synapses over several minutes appeared to be at a steady-state, thus not changing the overall size of synapses.

FIGURE 9.3. Lateral mobility of NMDA receptors.

FIGURE 9.3

Lateral mobility of NMDA receptors. (a) Agonist-evoked block of synaptic receptors. Whole-cell application (top panel and arrows in lower panel, for 1 sec) of NMDA in the presence of MK801 resulted in complete and irreversible block of the NMDA receptor-mediated (more...)

Synaptic NMDA receptors preferentially contain the NR2A subunit, whereas NR1/NR2B assemblies largely form extrasynaptic receptors [56] and should be sensitive to the NR2B-specific antagonist, ifenprodil [57]. With these criteria in mind, post-MK801 recovering EPSCs might be assumed to become sensitive to ifenprodil due to the import of extrasynaptic NR1/NR2B receptors. However, this did not occur, implying that synaptic receptors might be just as mobile as extrasynaptic receptors in that recovery of synaptic receptors could actually be occurring because of the import of other synaptic receptors as well as, or instead of, extrasynaptic receptors.

This study by Tovar and Westbrook [52] emphasized how the synaptic complement of NMDA receptors are very much more dynamic than previously realized. This result raises the possibility that the trafficking and targeting of functional forms of the NMDA receptor are realistic mechanisms for modulating synaptic strength. Clearly, a change in the number and type of synaptic NMDA receptor would alter the receptor mediated Ca2+ concentration in spines, with important implications for the activity of kinases and phosphatases mediating changes in synaptic strength.

9.8. SYNAPTIC INHIBITION: THE MOBILITY OF EXTRASYNAPTIC GABAA RECEPTORS

If the general organization of the inhibitory synapse is not dissimilar to that of the excitatory synapse, one might expect a similar degree of regulation of receptor traffic. Molecular scaffold proteins (such as gephyrin and other associated proteins like GABARAP, Raft, Collibistin and Plic1 [58,59]) are stabilized by the cytoskeleton, an association that ensures the sub-synaptic localization of glycine and GABAA receptors. However, much recent evidence from confocal microscopy studies has revealed that inhibitory synapses are also subject to rapid structural modifications [13]. In fact, the application of single particle and quantum dot tracking of metabotropic glutamate receptors, AMPA receptors, NMDA receptors and glycine receptors, within and outside the synapse, has established lateral mobility of these receptors as an increasingly important mechanism in the organization of the post-synaptic membrane [53,54,60,61].

We have recently studied the dynamics of functional GABAA receptors at synaptic and extrasynaptic loci in hippocampal neurons, specifically to establish whether these receptors are as mobile as other LGIC members. Unfortunately, unlike many of the other LGICs, the GABAA receptor does not have the benefit of an irreversible blocker of receptor function in its pharmacopoeia (such as MK-801 for the NMDA receptor or α-bungarotoxin for the nACh receptor). Nor does any evidence exist that the biophysical profile of a GABAA receptor can be sufficiently changed by the import or export of specific receptor subtypes into or out of the synapse in response to stimulation. Thus, functional studies of this nature have been hampered by the lack of an inherent electrophysiological tag. To overcome this condition, we engineered a silent mutation into the channel lining transmembrane domain of the α1 subunit of the receptor, at a location previously revealed by systematic cysteine scanning mutagenesis [62]. By mutating a hydrophobic residue in the ion channel region to a cysteine at the 2′ position (V257C), the application of the cysteine-modifying reagent MTSES to mutant recombinant α1V257Cβ1γ2 receptors was largely ineffective in the absence of GABA. However, after receptor activation by GABA, and presumably full exposure of the cysteine residue to MTSES, an irreversible 75% reduction in current amplitude was observed (Figure 9.4a) [62]. This difference in sensitivity to MTSES between open and closed GABA channels and its predictable use-dependence has formed the basis for a functional tag of α1-containing GABAA receptors in hippocampal neurons [63]. Transfection of neurons with the mutated subunit cDNA allowed the correct assembly and transport of the mutant subunit to appropriate locations frequented by wild-type α1 subunits. When these mutant receptors are located at synaptic sites, thus becoming subject to spontaneously-released GABA, they will also be susceptible to block by MTSES, whereas those extrasynaptic receptors not “seeing” released GABA would remain resistant. Under these conditions, the normal function of synaptic receptors (monitored as mIPSCs) is inhibited by up to 50% following MTSES exposure (Figure 9.4b). The reaction of MTSES with the channel cysteine is covalent and thus irreversible; therefore, any recovery of receptor function will come from either direct insertion into the synapse of unblocked receptor or from lateral diffusion of extrasynaptic receptors. Both scenarios are possible because MTSES is not membrane permeant and the absence of GABA means that intracellular pools of receptor will be unaffected by MTSES, and extrasynaptic receptors have not been exposed to the same concentrations of GABA as those at the synapse and thus remain unblocked. We can largely discount tonic levels of GABA because the experiments were performed on low-density cultures and only mIPSC events were recorded, so spillover of neurotransmitter was kept to a minimum. With this experimental paradigm, we were able to observe the very rapid recovery of blocked synaptic receptors within 10 to 20 min of the maximal MTSES effect (Figure 9.4b). The unblocked receptors appeared to move into the synapse, replacing blocked receptors by lateral diffusion because inhibitors of vesicular, exocytotic delivery (i.e., botulinum toxin B and N-ethylmaleimide) were unable to affect the basal delivery of GABAA receptors to the membrane from inside the cell during the timescale of the recovery. Moreover, blockade of all surface receptors in the neurone (synaptic plus extrasynaptic) by MTSES resulted in no recovery over a longer timescale, suggesting that surface receptor mobility is key to the recovery process.

FIGURE 9.4. Tracking synaptic GABAA receptor movements.

FIGURE 9.4

Tracking synaptic GABAA receptor movements. (a) Application of 10-mM MTSES to Xenopus oocytes injected with the α1V257Cβ1γ 2 subunits of the GABAA receptor show irreversible current blockade in the presence (left panel) but not (more...)

Taken overall, these observations indicate a very slow turnover rate of GABAA receptors by endo- and exocytotic processes, and would thus largely discount receptor delivery from within the cell as the recovery mechanism. The more likely explanation is that recovery of synaptic GABAA receptors results from lateral diffusion of unblocked receptors into the synapse, restoring the efficacy of synaptic transmission. This represents the first study using an electrophysiological tag to establish the dynamics of GABAA receptors in neurons.

9.9. PHOTORECEPTIVE POTASSIUM CHANNELS AS SWITCHES OF NEURONAL EXCITABILITY

The ion-channel pore belonging to fast-acting, ligand-gated receptors has proved a useful locus for functional receptor tags. This approach could also be applied to voltage-gated ion channels. An innovative variation on this theme has recently evolved for the Shaker K+ channel, which, like other members of this family, is susceptible to reversible channel block by quaternary ammonium ions such as tetraethylammonium (TEA). This channel has been tagged by building a novel “chemical gate” that controls the activation/inactivation of the channel regulated by δ light of particular wavelengths. The elegant generation of this so-called SPARK (synthetic photoisomerizable azobenzene-regulated K+, Figure 9.5a) channel involves the tethering of a synthetic maleimide-azobenzene derivative to an introduced cysteine residue at a known location in the S5 and S6 pore unit of the Shaker potassium channel [64,65]. This cysteine is sufficiently close to the ion channel that TEA, attached to one end of the tether, can bind to its site in the ion channel and block ion flow. In the case of the photoisomerizable azobenzene group, the wavelength of light determines whether the azo moiety adopts a cis or trans conformation consequently shortening or lengthening the tether and so either removing or allowing block of the ion channel by TEA (Figure 9.5b). Exposure to UV light (380 nm) induces the cis conformation that shortens the linker and effectively pulls the TEA moiety at its extremity away from the channel δ-preventing pore block (Figure 9.5b), whereas visible light (460 to 500 nm) allows revision of the molecule to the thermodynamically δ-favored trans conformation, which is longer by 7Å, re-establishing channel block. The tethered ligand thus acts as a light-activated chemical gate. Switching between these two states of block and unblock is achieved very rapidly. The protein modification required to permit attachment of the synthetic molecule introduces a cysteine into the protein to which a maleimide molecule at one end of the tether attaches in an irreversible manner. This mutation is functionally silent. Once attached, under visible light the tether passively blocks the K+ channel, ablating almost 1 nA of current in oocytes (Figure 9.5c).

FIGURE 9.5. Engineering membrane channel inactivation with light.

FIGURE 9.5

Engineering membrane channel inactivation with light. (a) Synthetic maleimide-azobenzene-TEA (MAL-AZO-QA) tether used in the modification of the Shaker K+ channel. The maliemide moiety attaches to a cysteine residue; the azobenzene moiety changes the (more...)

Following appropriate mutation of the SPARK channel to increase the neuronal resting conductance for K+ (by eliminating rapid inactivation) and subsequent expression in hippocampal neurons, spontaneous action potential activity was markedly reduced. The activation of the SPARK channels via the tethered TEA resulted in a general increase in neuronal activity within seconds of exposure to UV light (due to channel block, Figure 9.5d). Clearly, this technique could have a number of applications, not least in the programmable silencing of other functional ion channels or LGICs, whose membrane dynamics could subsequently be tracked. The convenient nature of the channel block during episodes of normal light (i.e., due to the favored trans configuration) would allow functional receptors to be exposed selectively under UV light.

9.10. DISADVANTAGES AND LIMITATIONS OF THE FUNCTIONAL TAG

Many of the limitations of the functional tag assay are also applicable to biochemical or optical methods, though any modification of a protein that might affect its normal function (i.e., current flux) should be thoroughly assessed in the isolation of a recombinant system prior to its use in a native environment. In particular the tag must be functionally silent until activated or bound by a ligand. Thus, one needs to check ion-channel gating in response to agonist, whether any alteration to agonist potency is found, or if unusual rectification properties are present after receptor activation. In addition, and again common to optical and biochemical tracking methods, ensuring that the use of a functional tag does not introduce an epitope that could inadvertently trigger endocytosis or affect interactions with accessory proteins is necessary, for example, by disrupting a receptor’s phosphorylation state, anchoring or trafficking. All these aspects can be checked in control experiments. For these reasons, if a receptor has an inherent functional tag, then this should be exploited in preference to engineered tags by mutagenesis. Clearly, if an epitope must be introduced, the affects of expressing a protein in a neurone must then also be considered. Many transfection protocols, especially those using viral vectors, have become very efficient even in primary cell cultures. The consequence of this efficiency is that the cell could be overwhelmed with exogenous DNA products, causing the basal equilibrium of protein manufacture to be upset. Under these conditions, certain LGICs might be inappropriately targeted or trafficked, or others might be under-represented in the cell membrane. All these considerations must be taken into account when interpreting the data.

9.11. CONCLUSION

Functional tagging of receptor proteins offers a unique method of tracking receptor movement. It has one overriding advantage in that, unequivocally, one is measuring the movement of functional surface receptors by virtue of the fact that they can be activated and pass current. The method does not need to rely on co-localization with other proteins to assume importance at synapses, nor is inferring that they are functional receptors necessary. In the future, use of more innovative silent tags will allow the resolution of receptor translocation in the surface membrane to become even more accurate, and with more specific inhibitors of exo- and endocytotic processes, we will gain a better understanding of the life-cycle of important neurotransmitter receptors in the central nervous system.

REFERENCES

1.
Betz WJ, Mao F, Bewick GS. Activity-dependent fluorescent staining and destaining of living vertebrate motor nerve terminals. J Neurosci. 1992;12:363–375. [PubMed: 1371312]
2.
Betz WJ, Mao F, Smith CB. Imaging exocytosis and endocytosis. Curr Opin Neurobiol. 1996;6:365–371. [PubMed: 8794083]
3.
Kannenberg K, Sieghart W, Reuter H. Clusters of GABAA receptors on cultured hippocampal cells correlate only partially with functional synapses. Eur J Neurosci. 1999;11(4):1256–1264. [PubMed: 10103120]
4.
Passafaro M, Piech V, Sheng M. Subunit-specific temporal and spatial patterns of AMPA receptor exocytosis in hippocampal neurons. Nat Neurosci. 2001;4:917–926. [PubMed: 11528423]
5.
Miesenbock G, De Angelis DA, Rothman JE. Visualizing secretion and synaptic transmission with pH-sensitive green fluorescent proteins. Nature. 1998;394:192–195. [PubMed: 9671304]
6.
Ashby MC, De La Rue SA, Ralph GS, Uney J, Collingridge GL, Henley JM. Removal of AMPA receptors (AMPARs) from synapses is preceded by transient endocytosis of extrasynaptic AMPARs. J Neurosci. 2004;24:5172–5176. [PMC free article: PMC3309030] [PubMed: 15175386]
7.
Zhang J, Campbell RE, Ting AY, Tsien RY. Creating new fluorescent probes for cell biology. Nat Rev Mol Cell Biol. 2002;3:906–918. [PubMed: 12461557]
8.
Gaietta G, Deerinck TJ, Adams SR, Bouwer J, Tour O, Laird DW, Sosinsky GE, Tsien RY, Ellisman MH. Multicolor and electron microscopic imaging of connexin trafficking. Science. 2002;296:503–507. [PubMed: 11964472]
9.
Ju W, Morishita W, Tsui J, Gaietta G, Deerinck TJ, Adams SR, Garner CC, Tsien RY, Ellisman MH, Malenka RC. Activity-dependent regulation of dendritic synthesis and trafficking of AMPA receptors. Nature Neurosci. 2004;7:244–253. [PubMed: 14770185]
10.
Sekine-Aizawa Y, Huganir RL. Imaging of receptor trafficking by using alpha bungarotoxin-binding-site-tagged receptors. Proc Natl Acad Sci USA. 2004;101(49):17114–17119. [PMC free article: PMC534416] [PubMed: 15563595]
11.
Weiss RE, Roberts WM, Stuhmer W, Almers W. Mobility of voltage-dependent ion channels and lectin receptors in the sarcolema of frog skeletal muscle. J Gen Physiol. 1986;87:955–983. [PMC free article: PMC2215866] [PubMed: 2425044]
12.
Young SH, Poo M.-M. Rapid lateral diffusion of extra-junctional acetylcholine receptors in the developing muscle membrane of Xenopus tadpole. J Neurosci. 1983;3:225–231. [PubMed: 6822857]
13.
Choquet D, Triller A. The role of receptor diffusion in the organization of the postsynaptic membrane. Nat Rev Neurosci. 2003;4(4):251–265. [PubMed: 12671642]
14.
Farrant M, Nusser Z. Variations on an inhibitory theme: Phasic and tonic activation of GABAA receptors. Nat Rev Neurosci. 2005;6(3):215–229. [PubMed: 15738957]
15.
Mody I. Aspects of the homeostaic plasticity of GABAA receptor-mediated inhibition. J Physiol. 2005;562(1):37–46. [PMC free article: PMC1665492] [PubMed: 15528237]
16.
Semyanov A, Walker MC, Kullmann DM, Silver RA. Tonically active GABAA receptors modulating gain and maintaining the tone. Trends Neurosci. 2004;27(5):262–269. [PubMed: 15111008]
17.
Poo M.-M, Cone RA. Lateral diffusion of rhodopsin in the photoreceptor membrane. Nature. 1974;247:438–441. [PubMed: 4818543]
18.
Axelrod D, Ravdin P, Koppel DE, Schlessinger J, Webb WW, Elson EL, Podleski TR. Lateral motion of fluorescently labeled acetylcholine receptors in membranes of developing muscle fibers. Proc Natl Acad Sci USA. 1976;73(12):4594–4598. [PMC free article: PMC431558] [PubMed: 1070010]
19.
Almers W, Stirling C. Distribution of transport proteins over animal cell membranes. J Memb Biol. 1984;77(3):169–186. [PubMed: 6321741]
20.
Kim E, Sheng M. PDZ domain proteins of synapses. Nat Rev Neurosci. 2004;5(10):771–781. [PubMed: 15378037]
21.
Li Z, Sheng M. Some assembly required: The development of neuronal synapses. Nat Rev Mol Cell Biol. 2003;4(11):833–841. [PubMed: 14625534]
22.
Sheng M. Molecular organization of the postsynaptic specialization. Proc Natl Acad Sci USA. 2001;98(13):7058–7061. [PMC free article: PMC34622] [PubMed: 11416187]
23.
Benke TA, Luthi A, Isaac JT, Collingridge GL. Modulation of AMPA receptor unitary conductance by synaptic activity. Nature. 1998;393:793–797. [PubMed: 9655394]
24.
Roche KW, O’Brien RJ, Mammen AL, Bernhardt J, Huganir RL. Characterization of multiple phosphorylation sites on the AMPA receptor GluR1 subunit. Neuron. 1996;16:1179–1188. [PubMed: 8663994]
25.
Hayashi Y, Ishida A, Katagiri H, Mishina M, Fujisawa H, Manabe T, Takahashi T. Calcium- and calmodulin-dependent phosphorylation of AMPA type glutamate receptor subunits by endogenous protein kinases in the post-synaptic density. Mol Brain Res. 1997;46(1-2):338–342. [PubMed: 9191113]
26.
Barria A, Muller D, Derkach V, Griffith LC, Soderling TR. Regulatory phosphorylation of AMPA-type glutamate receptors by CaM-KII during long-term potentiation. Science. 1997;276(5321):2042–2045. [PubMed: 9197267]
27.
Derkach V, Barria A, Soderling TR. Ca2+/calmodulin-kinase II enhances channel conductance of alpha-amino-3-hydroxy-5-methyl-4-isoxazolepropionate type glutamate receptors. Proc Natl Acad Sci USA. 1999;96(6):3269–3274. [PMC free article: PMC15931] [PubMed: 10077673]
28.
Petralia RS, Esteban JA, Wang YX, Partridge JG, Zhao HM, Wenthold RJ, Malinow R. Selective acquisition of AMPA receptors over postnatal development suggests a molecular basis for silent synapses. Nat Neurosci. 1999;2:31–36. [PubMed: 10195177]
29.
Lissin DV, Gomperts SN, Carroll RC, Christine CW, Kalman D, Kitamura M, Hardy S, Nicoll RA, Malenka R, von Zastrow M. Activity differentially regulates the surface expression of synaptic AMPA and NMDA glutamate receptors. Proc Natl Acad Sci USA. 1998;95(12):7097–7102. [PMC free article: PMC22752] [PubMed: 9618545]
30.
O’Brien RJ, Kamboj S, Ehlers MD, Rosen KR, Fischbach GD, Huganir RL. Activity-dependent modulation of synaptic AMPA receptor accumulation. Neuron. 1998;21:1067–1078. [PubMed: 9856462]
31.
Turrigiano GG, Leslie KR, Desai NS, Rutherford LC, Nelson SB. Activity-dependent scaling of quantal amplitude in neocortical neurons. Nature. 1998;391:892–896. [PubMed: 9495341]
32.
Liu SJ, Cull-Candy SG. Synaptic activity at calcium-permeable AMPA receptors induces a switch in receptor subtype. Nature. 2000;405:454–458. [PubMed: 10839540]
33.
Rozov A, Burnashev N. Polyamine-dependent facilitation of postsynaptic AMPA receptors counteracts paired-pulse depression. Nature. 1999;401:594–598. [PubMed: 10524627]
34.
Bowie D, Mayer ML. Inward rectification of both AMPA and kainate subtype glutamate receptors generated by polyamine-mediated ion channel block. Neuron. 1995;15:453–462. [PubMed: 7646897]
35.
Kamboj SK, Swanson GT, Cull-Candy SG. Intracellular spermine confers rectification on rat calcium-permeable AMPA and kainate receptors. J Physiol. 1995;486(2):297–303. [PMC free article: PMC1156521] [PubMed: 7473197]
36.
Koh DS, Burnashev N, Jonas P. Block of native Ca2+-permeable AMPA receptors in rat brain by intracellular polyamines generates double rectification. J Physiol. 1995;486(2):305–312. [PMC free article: PMC1156522] [PubMed: 7473198]
37.
Sommer B, Kohler M, Sprengel R, Seeburg PH. RNA editing in brain controls a determinant of ion flow in glutamate-gated channels. Cell. 1991;67(1):11–19. [PubMed: 1717158]
38.
Nusser Z, Lujan R, Laube G, Roberts JD, Molnar E, Somogyi P. Cell type and pathway dependence of synaptic AMPA receptor number and variability in the hippocampus. Neuron. 1998;21:545–559. [PubMed: 9768841]
39.
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: 10409387]
40.
Zhu JJ, Malinow R. Acute versus chronic NMDA receptor blockade and synaptic AMPA receptor delivery. Nat Neurosci. 2002;5:513–514. [PubMed: 11967548]
41.
Hayashi Y, Shi SH, Esteban JA, Piccini A, Poncer JC, Malinow R. Driving AMPA receptors into synapses by LTP and CaMKII: Requirement for GluR1 and PDZ domain interaction. Science. 2000;287(5461):2262–2267. [PubMed: 10731148]
42.
Luscher C, Xia H, Beattie EC, Carroll RC, von Zastrow M, Malenka RC, Nicoll RA. Role of AMPA receptor cycling in synaptic transmission and plasticity. Neuron. 1999;24:649–658. [PubMed: 10595516]
43.
Allison DW, Gelfand VI, Spector I, Craig AM. Role of actin in anchoring postsynaptic receptors in cultured hippocampal neurons: Differential attachment of NMDA versus AMPA receptors. J Neurosci. 1998;18:2423–2436. [PubMed: 9502803]
44.
Kennedy MB. Signal-processing machines at the postsynaptic density. Science. 2000;290(5492):750–754. [PubMed: 11052931]
45.
Huh KH, Wenthold RJ. Turnover analysis of glutamate receptors identifies a rapidly degraded pool of the N-methyl-D-aspartate receptor subunit, NR1, in cultured cerebellar granule cells. J Biol Chem. 1999;274(1):151–157. [PubMed: 9867823]
46.
Malenka RC, Nicoll RA. Long-term potentiation: A decade of progress? Science. 1999;285(5435):1870–1874. [PubMed: 10489359]
47.
Mori H, Mishina M. Structure and function of the NMDA receptor channel. Neuropharmacology. 1995;34(10):1219–1237. [PubMed: 8570021]
48.
Carroll RC, Zukin RS. NMDA-receptor trafficking and targeting: Implications for synaptic transmission and plasticity. Trends Neurosci. 2002;25(11):571–577. [PubMed: 12392932]
49.
Wenthold RJ, Prybylowski K, Standley S, Sans N, Petralia RS. Trafficking of NMDA receptors. Ann Rev Pharm Toxicol. 2003;43:335–358. [PubMed: 12540744]
50.
Wenthold RJ, Sans N, Standley S, Prybylowski K, Petralia RS. Early events in the trafficking of N-methyl-D-aspartate (NMDA) receptors. Biochem Soc Trans. 2003;31(4):885–888. [PubMed: 12887327]
51.
Lan JY, Skeberdis VA, Jover T, Grooms SY, Lin Y, Araneda RC, Zheng X, Bennett MV, Zukin RS. Protein kinase C modulates NMDA receptor trafficking and gating. Nat Neurosci. 2001;4:382–390. [PubMed: 11276228]
52.
Tovar KR, Westbrook GL. Mobile NMDA receptors at hippocampal synapses. Neuron. 2002;34:255–264. [PubMed: 11970867]
53.
Groc L, Heine M, Cognet L, Brickley K, Stephenson FA, Lounis B, Choquet D. Differential activity-dependent regulation of the lateral mobilities of AMPA and NMDA receptors. Nat Neurosci. 2004;7:695–696. [PubMed: 15208630]
54.
Borgdorff AJ, Choquet D. Regulation of AMPA receptor lateral movements. Nature. 2002;417(6889):649–653. [PubMed: 12050666]
55.
Meier J, Vannier C, Serge A, Triller A, Choquet D. Fast and reversible trapping of surface glycine receptors by gephyrin. Nat Neurosci. 2001;4(3):253–260. [PubMed: 11224541]
56.
Tovar KR, Westbrook GL. The incorporation of NMDA receptors with a distinct subunit composition at nascent hippocampal synapses in vitro. J Neurosci. 1999;19:4180–4188. [PubMed: 10234045]
57.
Williams K. Ifenprodil discriminates subtypes of the N-methyl-D-aspartate receptor: Selectivity and mechanisms at recombinant heteromeric receptors. Mol Pharm. 1993;44(4):851–859. [PubMed: 7901753]
58.
Luscher B, Keller CA. Regulation of GABAA receptor trafficking, channel activity, and functional plasticity of inhibitory synapses. Pharm Ther. 2004;102(3):195–221. [PubMed: 15246246]
59.
Moss SJ, Smart TG. Constructing inhibitory synapses. Nat Rev Neurosci. 2001;2(4):240–250. [PubMed: 11283747]
60.
Tardin C, Cognet L, Bats C, Lounis B, Choquet D. Direct imaging of lateral movements of AMPA receptors inside synapses. EMBO J. 2003;22(18):4656–4665. [PMC free article: PMC212729] [PubMed: 12970178]
61.
Serge A, Fourgeaud L, Hemar A, Choquet D. Receptor activation and homer differentially control the lateral mobility of metabotropic glutamate receptor 5 in the neuronal membrane. J Neurosci. 2002;22:3910–3920. [PubMed: 12019310]
62.
Xu M, Akabas MH. Identification of channel-lining residues in the M2 membrane-spanning segment of the GABAA receptor alpha1 subunit. J Gen Physiol. 1996;107(2):195–205. [PMC free article: PMC2219269] [PubMed: 8833341]
63.
Thomas P, Mortensen M, Hosie AM, Smart TG. Dynamic mobility of functional GABAA receptors at inhibiting synapses. Nab Neurosci. 8:889–8997. [PubMed: 15951809]
64.
Blaustein RO, Cole PA, Williams C, Miller C. Tethered blockers as molecular “tape measures” for a voltage-gated K+ channel. Nat Struct Biol. 2000;7(4):309–311. [PubMed: 10742176]
65.
Banghart M, Borges K, Isacoff E, Trauner D, Kramer RH. Light-activated ion channels for remote control of neuronal firing. Nat Neurosci. 2004;7:1381–1386. [PMC free article: PMC1447674] [PubMed: 15558062]
Copyright © 2006, Taylor & Francis Group, LLC.
Bookshelf ID: NBK2550PMID: 21204475

Views

  • PubReader
  • Print View
  • Cite this Page

Other titles in this collection

Related information

  • PMC
    PubMed Central citations
  • PubMed
    Links to PubMed

Similar articles in PubMed

See reviews...See all...

Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...