Qual Journals
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Tardin, Choquet et al. (2003). Direct imaging of lateral movements of AMPA receptors inside synapses. The EMBO Journal, 22(18)
- Statements
Trafficking of AMPAR in and out of synapses is crucial for synaptic plasticity. Protocols that induce plasticity of synaptic transmission in culture result in changes of AMPAR concentration at synapses and are thought to mimic at the molecular level the processes of LTP and LTD.
Membrane trafficking may occur outside of the synapse and accumulate at the PSD after a short delay (Passafaro et al. 2001). Altogether, a unified picture of the postsynaptic density could be one where receptors are immobilized for transient periods of time related to the receptor-scaffold affinity. This could also be true of NMDA receptors (Tovar and Westbrook, 2002).
- Findings
Application of glutamate increased the diffusion rate of GluR2-containting AMPAR whereas a protocol designed to induce calcium influx (stimulation of NMDAR with glycine, glutamate) reduced the percentage of diffusible AMPARs at the PSD. Bath application of 100 uM glutamate caused an 85% increase in AMPAR endocytosis within 15 min (corresponding to a 22% drop in total membrane expression). Conversely , the calcium influx protocol (20 uM biccuculine, 1 uM strychnine, 200 uM glycine) caused a 59% increase in AMPAR membrane expression. Glutamate caused a 55% increase in AMPAR diffusion within synapses, but did not change diffusion outside synapses. Furthermore, glutamate decreased the number of completely immobile AMPARs by 30%. Interestingly, Glutamate causes endocytosis of AMPARs, and internal AMPARs are immobile. Therefore it seems like glutamate may be causing a general endocytotic episode at non-synaptic AMPARs, perhaps not even at the synapse that received the glutamate application. In a parallel effect, it was found that blocking calcium with BAPTA increased the % of mobile AMPARs. Newly inserted receptors were found to be initially diffusive and then stabilized at synaptic sites. In summary, they found that bath application of glutamate induces rapid depletion of AMPARs from PSDs increases synaptic diffusion rate, decreases % of completely immobile receptors, increases proportion of receptors in the area surrounding the synapse (juxtasynaptic region). Activation of NMDARs results in increased surface expression of AMPARs -- in the first few minutes there is mainly a decrease in the proportion of immobile synaptic receptors, but after 40 min, both diffusion rates and percentages of immobile synaptic receptors are back to control values and the proportion of juxtasynaptic receptors is decreased. This observation relates to the fate of newly exocytosed AMPARs: using cleavable extracellular tags, it was observed that at early times after exocytosis, new GluR1 containing AMPARs are diffusively distributed along dendrites. This is followed by their lateral translocation and accumulation into synapses (Passafaro et al., 2001). GluR2 subunits were addressed directly at synapses. In our experiments, we followed the movement of native GluR2 containing AMPARs, where the data suggests that at the level of synapses themselves, newly added receptors are initially diffusive and then stabilize over time.
AMPA receptors that lack edited GluA2 subunits have high single channel conductance, are permeable to Ca2+, are blocked by polyamines causing inward rectification at depolarized potentials.
- 100 uM glutamate - within 15 min
- 85% increase in AMPAR endocytosis
- 22% drop in total membrane expression
- 55% increase in AMPAR diffusion rate within synapses
- 0% increase in AMPAR diffusion rate outside synapses
- 30% decrease in completely immobile AMPAR at PSD
--
- Start: 100 AMPARs in PSD
- Usual endocytosis rate: -0.25% / min
- Add: 100 uM glutamate
- New endocytosis rate: -1.5% / min
- Time: 15 min
- Final: 77.5 AMPARs
- calcium influx protocol (20 uM biccuculine, 1 uM strychnine, 200 uM glycine)
- 59% increase in AMPAR expression
--
- Start: 100 AMPARs in PSD
- Usual endocytosis rate: -0.25% / min
- Add: NMDA antagonists above
- New exocytosis rate: +4% / min
- Time: 15 min
- Final: 160 AMPARs
Retrieval-specific endocytosis of GluA2-AMPARs underlies adaptive reconsolidation of contextual fear
Rao-Ruiz, Spijker, et al.
A consolidated memory returns to a transient destabilized state shortly after reactivation, necessitating a dynamic time-dependent process of reconsolidation to persist further. During this reconstruction, a memory is labile and subject to change. In general, a memory-recall causes internalization of AMPAR at activate synapses for ~2 hours. Then AMPAR repopulate these synapses and return to baseline levels, and sometimes even higher levels.
- GROUPS
- NS: No Shock
- US: Shock
- NR: No Retrieval (24h later)
- R: Retrieval (24h later)
Tissue collected 1 h after retrieval for western blot and ephys
RESULTS
Down-regulation of all AMPAR subtypes and smaller mEPSC amplitudes 1 h after retrieval. Increase at 7 h.
Start at 100% baseline expression
- Post-Recall: 1 h 4 h 7 h
- GluR1 70% 100% 100%
- GluR2 85% 85% 130%
- GluR3 50% 50% 100%
Blocking GluR2 internalization with 3Y peptide (3A is a control peptide) 1 h before or 1 h after recall prevented a subsequent GluR2 increase at 7 hours. The narrow decay time found at 7 hours in 3A controls suggests that synapses are composed more of GluR2 than GluR1 or GluR3 receptors which are permeable to calcium. Furthermore, 3Y increased fear memory when tested 2 h (RT2) after a recall event (RT1) or 24 h (RT3) after recall.
- Start at 100% baseline expression
- Post-Recall: 1 h 4 h 7 h
- GluR1 70% 100% 100%
- GluR2 85% 85% 130%
- GluR3 50% 50% 100%
MAKING QUANTUM DOTS
To create a quantum dot that will tag AMPA receptors choose a quantum dot conjugated to an IgG antibody that also has a fragment antigen-binding region (aka F(ab’)2 or Fab fragment, the part of an antibody that binds antigens). This entire construct, QD-IgG-Fab, is your secondary antibody construct and the IgG and Fab are usually from different host animals. This is then incubated with primary antibody fragment (Fab) that is specific to the protein of interest, in this case AMPA receptors.
Single-Nanoparticle Tracking.
A total of 1 μL of 655-nm Quantum dots (Qdots) conjugated (see also antibodies) to goat (Fab‘)2 anti-mouse IgG (Invitrogen) were incubated with 1 μL Fab anti-GluA2 in 7 μL PBS for 20 min at room temperature. Nonspecific binding was blocked by adding 1 μL of 10% casein stock solution for 15 min (Vector Laboratories), and this solution was kept at 4 °C throughout the experiment. Neurons were incubated for 10 min at 37 °C in 1 mL culture medium containing 1 μL of the anti-GluA2-coated Qdot solution, then rinsed and mounted in an aluminum chamber containing Tyrode solution (30 mM glucose, 120 mM NaCl, 5 mM KCl, 2 mM MgCl2, 2 mM CaCl2, 25 mM Hepes) on a Nikon microscope (NIKON Eclipse TE2000-U) thermostated to 37 °C using an air blower (World Precision Instruments) and an objective heater (Bioptechs). Single Qdots were detected through a100 × 1.4 N.A. oil immersion objective, using a 100-W mercury lamp and appropriate excitation/emission filters (Chroma). Sequences of 50 s, corresponding to stacks of 1,000 images with an integration time of 50 ms, were acquired using a CCD camera (Quantem; Roper Scientific). For each coverslip, Qdots were followed on randomly selected dendritic regions (size 20 × 20 μ m) for up to 20 min. Two coverslips of each condition were processed for a total of 3 – 4 neuronal cultures. We checked for specificity of Qdot binding by preparing primary hippocampal cultures from GluA2 KO mice dissected at p0. Anti – GluA2- conjugated Qdots bound minimally to cultures from GluA2 KO compared with those prepared from wild-type littermates.
Antibodies and drugs.
The antibody to the N-terminal epitope of the GluR1 subunit was described previously46. We used a commercial antibody to an N-terminal epitope of the GluR2 subunit to detect GluR2 (BD Pharmigen). AMPA receptor labeling and synaptic live staining. Quantum dot 655 goat F(ab')2 antibody to rabbit IgG conjugate (H+L) highly cross-absorbed and quantum dot 655 goat F(ab')2 antibody to mouse IgG conjugate (H+L) highly cross-absorbed were obtained from Quantum Dot (Invitrogen). Receptors were stained using quantum dots pre-coated with antibody to GluR1 or monoclonal antibody to GFP. Quantum dots (0.1 M) were incubated with 1 g of antibody in 10 l of phosphate-buffered saline (PBS) for 15–30 min. Unspecific binding was blocked by adding casein (Vector Laboratories) to the pre-coated quantum dot 15 min before use. Neurons were incubated 5–10 min at 37 °C in culture medium with pre-coated quantum dots (final dilution of 0.1–0.01 nM). The incubation was followed by four washing steps of 30 s each. All incubations and washes were performed in pre-warmed extracellular HEPES-buffered solution (see below).
Single molecule optical microscopy.
Cells were imaged at 35–37 °C in an open chamber mounted onto an inverted microscope (IX70 Olympus) equipped with a 60 (NA = 1.35, Olympus) or 100 objective (NA = 1.3, Olympus). Quantum dots and Homer1C-DsRed were detected using a xenon lamp (excitation filter HQ500/20X (Chroma), Mitotrack 560RDF55 (Omega)) and appropriate emission filters (HQ560/80M (Chroma Technology), 655WB20 (Omega Optical)). Fluorescent images from quantum dots were acquired with an integration time of 33 ms with up to 2,000 consecutive frames. Signals were recorded with a back-illuminated thinned CCD camera (Cascade 512BFT, Roper Scientific).
Quantum dot–labeled GluR1 receptors were monitored on randomly selected dendritic regions for up to 20 min of total experimental time. Recording of the synaptic marker over time revealed that the mobility of synapses was much slower in comparison to the mobility of the receptors (data not shown). Mobility of synapses themselves did not seem to affect our location method. Acquisition of the synaptic labeling before and after quantum dot recording as well as quantum dots fixed on the cover slip allowed us to compensate mechanical drifts of the stage, which would have lead to a false interpretation of receptor location.
Receptor tracking and analysis.
The tracking of single quantum dots was performed with homemade software based on Mathlab (Mathworks). Single quantum dots were identified by their blinking fluorescent emission and their diffraction-limited signals. Owning to the random blinking events of the quantum dots, the trajectory of a quantum dot–tagged receptor could not be tracked continuously. Subtrajectories of the same receptor were reconnected when the positions before and after the dark period were compatible with borders set for maximal position changes between consecutive frames and blinking rates. The values were determined empirically: 1–2 pixels for maximal position change between two frames and maximal dark periods of 25 frames. Mean square displacement curves were calculated for reconnected trajectories of at least 100 frames. Diffusion coefficients were calculated by a linear fit of the first four points of the msd plots versus time. The resolution limit for diffusion was 0.001 m2 s-1, as determined by msd calculations of fixed quantum dots. The resolution precision was 40 nm. Dwell times of individual receptors given in the results were measured from trajectories in which the entry and exit from the compartments could be identified. Synaptic or ECM compartments were identified by homer1c-DsRed expression or HABP staining, respectively. Pixels assigned to synapses or ECM were defined as a set of connected pixels obtained using two-dimensional object segmentation by wavelet transformation47
Electrophysiology.
The extra cellular medium contained 145 mM NaCl, 2.5 mM KCl, 2 mM MgCl2, 2 mM CaCl2, 10 mM HEPES and 10 mM D-glucose (pH 7.4). To block GABAA receptors, we added 50 M picrotoxin to the extra cellular medium. The bath temperature was kept at 33–35 °C. Borosilicate pipettes were used to produce patch electrodes with resistances of 3–5 M . A standard pipette solution was used to characterize neuronal properties in voltage and current-clamp conditions during development (Supplementary Fig. 7) and contained 140 mM potassium gluconate, 2 mM MgCl2, 4 mM NaATP, 0.1 mM EGTA, 10 mM HEPES, 10 mM phosphocreatine, 0.4 mM GTP (pH 7.25). To record mEPSC and to decrease space-clamp difficulties, we used another recording solution 125 mM CH3CsSO3, 2 mM MgCl2, 1 mM CaCl2, 4 mM NaATP, 10 mM EGTA, 10 mM HEPES and 0.4 mM GTP (pH 7.25). Recordings in voltage and current clamp mode were performed with an EPC10 double patch-clamp amplifier (HEKA Electronics). Data were acquired and stored using Pulse-Pulse fit software version 8.62 (HEKA Electronics, Lambrecht, Germany) and analyzed with IGOR (WaveMetrics) and GraphPad Prism software. Spontaneous events were analyzed by Minianalysis (Synaptosoft). Local activation of receptors was performed by iontophoresis of glutamate using an amplifier from NPI Electronics. Pipettes for iontophoretic stimulation had resistances between 40–60 M when filled with 150 mM sodium glutamate (pH 7.4). A small retaining current was needed to keep glutamate inside the pipette (usually between 10–50 nA). Current pulses between 30 and 600 nA and 1–2 ms duration were required to evoke AMPAR-mediated currents between amplitudes of 30–600 pA under control conditions.
Outside-out recordings.
Outside-out patches were pulled from 14–21 DIV neurons. Internal solution contained 130 mM CsCl, 2 mM MgCl2, 10 mM EGTA, 10 mM HEPES and 4 mM Na2ATP. Pipette resistance was 3.5–4.5 M . After patch formation, the pipette was placed under the flow of a theta application pipette containing HEPES-buffered solution in one line and HEPES-buffered solution, 10 mM glutamate and 20 mM sucrose in the other line to clearly visualize the interface between solutions. The application pipette was immerged in the bath and heated to 37 °C for at least 1 cm. It is thus assumed that solutions were close to that temperature. Fast application was achieved with a piezo-electric manipulator (Burleigh). After the recording, the application was controlled by measuring the junction current between the two solutions. To measure recovery from desensitization, it is important to verify that 1-ms applications effectively saturate receptors. We verified this by measuring the amplitude of the currents evoked by 1- or 100-ms applications (Supplementary Fig. 6). If the former was less than 80% of the latter, the recording was discarded; on average, current amplitudes were 497.2 160 pA (n = 10) for control and 334.3 143 pA (n = 10) for treated neurons, and the amplitude ratios (1 ms/100 ms) were 0.9 0.03 and 0.91 0.05 for control and treated neurons, respectively.
FRAP.
Transfected neurons (21–30 DIV) were placed on the heated stage (37 °C) of an inverted microscope (Leica CTR 6500) and continually perfused with preheated (37 °C) extracellular solution (composition as described above). For low-pH solution, HEPES was replaced by MOPS and adjusted to pH 5.5. To test the population of surface pHGFP-GluR1–containing AMPARs of a particular cell, we used a gravity-driven rapid solution exchange using a theta-glass electrode containing low-pH solution in one channel and NH4Cl (50 mM) in the other channel to determine the ratio between the fluorescent intensities48. Fluorescence was excited using a monochromator (Cairn) controlled by Metamorph software (Universal Imaging). To photobleach locally, we used a sapphire laser 488-20 (Coherent) at 30% power to avoid photodamage. The laser was coupled to the microscope via a galvometric mirror (Roper Scientific), which allowed us to photobleach several regions in a short time window. Recovery from photobleaching was monitored by consecutive acquisition at a 10-Hz acquisition rate. Recovery curves were corrected for continuous photobleaching and background noise as described elsewere49.
FLIP.
For FLIP experiments, the laser beam was parked at the dendritic shaft at a power of 10% and a additional 75% intensity filter was used to avoid photodamage in the continuous bleached region. Continuous laser illumination was interrupted during image acquisition at a frequency of 0.2 Hz. Control of surface expression and experimental conditions were similar to those for the FRAP experiments described above.