This work was supported by NIA/NIH grants R01AG070255 and R01AG073133 to AL. We also thank University of California Irvine Alzheimer&#39;s disease research center (UCI-ADRC) for providing the human tissue shown in this manuscript. The UCI-ADRC is funded by NIH/NIA grant P30 AG066519.

All research is performed in compliance with institutional guidelines and approved by the institutional Animal Care and Use Committee of the University of California Irvine (IACUC-1998-1388) and the University of Texas Medical Branch (IACUC-1803024). Temporal cortex from a non-Alzheimer&#39;s disease (AD) brain (female, 74 years old, postmortem interval 2.8 h) and an AD-brain (female, 74 years old, postmortem interval 4.5 h) were provided by the University of California Irvine Alzheimer&#39;s disease research center (UCI-ADRC). Informed consent for brain donation was obtained by UCI-ADRC.
NOTE: Unfixed human brain tissue should be treated as a source of blood borne pathogens (BBP). Accordingly, BBP training is needed prior to start experiments. This protocol is performed in a biosafety level 2 (BSL2) laboratory under BSL2 requirements. Guidelines and precautions within the laboratory include: no food or drinks allowed in the laboratory, good laboratory practices must be followed, personal protection equipment (gloves, gown, no open-toed shoes) is required, and the door must be closed at all times.
1. Xenopus oocyte microinjection preparation
<ol>
	<li>To make injection needles, pull 3.5 inches borosilicate tubes using a micropipette puller. Once the microinjection needles are pulled, use a microscope and a razor blade to cut off enough of the tip of the needle so that microinjection can be performed (usually between 2-3 mm). 
	NOTE: The length of the needle&#39;s tip to cut can vary.</li>
	<li>Prepare 1x Barth&#39;s solution by adding 200 mL of Barth&#39;s Stock Solution (5x) to 800 mL of distilled water. Fill a 24-well, flat bottom tissue culture plate with 18 &#176;C 1x Barth&#39;s solution.
	<ol>
		<li>Prepare 5x Barth&#39;s stock solution as follows. In 1 L of distilled water, add 25.71 g of NaCl (sodium chloride), 0.372 g of KCl (potassium chloride), 0.301 g of CaCl2 (calcium dichloride), 0.389 g of Ca(NO3)2(4H2O) (calcium nitrate tetrahydrate), 1.01 g of MgSO4(7H2O) (magnesium sulfate heptahydrate), 1.008 g of NaHCO3 (sodium bicarbonate), and 11.91 g HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid).</li>
	</ol>
	</li>
	<li>Isolate Xenopus oocytes using the protocols described in18,19 and using 2x magnified stereoscope, selecting healthy looking, stage V-VI oocytes. Place about 10 oocytes per well and use one of the wells to house non-injected oocytes to be used as control cells when performing recordings.</li>
	<li>Retrieve a small Petri dish, 1 cm tall and 6 cm in diameter, and place Nylon mesh inside so that it covers the bottom of the dish. Then, pour 15 mL of 1x Barth&#39;s solution to be used for performing microinjections of the oocytes.</li>
	<li>Place a nanoinjector along the midline of the microscope platform. Turn the magnet to the Off position and slowly move the nanoinjector to the desired position while supporting it carefully. When the nanoinjector is properly positioned, turn the magnet back to the fully On position and make sure that it is secure.</li>
	<li>For a sample size of 50.6 nL, use the following settings on the side of the nanoinjector: 1 is D (down), 2 is D, 3 is U (up), 4 is D, and 5 is U. Position a piece of self-sealing thermoplastic, or microcentrifuge tube cap, on the microscope stand, aligning it with the trajectory of the nanoinjector. 
	NOTE: 50 nL is near the maximum amount of injected material that the cytoplasm can withstand21.</li>
	<li>Fill up an insulin syringe about half its volume with mineral oil (about 0.5 mL/cc). Using this syringe, fill the microinjection needle with mineral oil. Ensure that a visible bead of oil is seen at the tip of the needle.</li>
	<li>Expose the nanoinjector plunger to a minimum of 1-2 cm. Do this by holding down the EMPTY button until the plunger is visible at desired length. Once the plunger is exposed, slowly and carefully place the mineral oil filled microinjection glass needle onto the nanoinjector plunger, and make sure that it fits well through the black resistance O-ring and stops at the white resistance ring. Ensure not to bend the nanoinjector plunger in the process.</li>
	<li>Once the microinjection glass needle is secure and in place, press EMPTY to void any potential air bubbles. Gently, with a paper tissue, clean off the excess mineral oil.</li>
</ol>
2. Loading the sample
<ol>
	<li>Prepare synaptosomal enriched preparations (samples) from the human brain region of interest as described in10,11,12,13,14,15, and store them in aliquots of about 5 &#956;L at -80&#160;&#176;C until the moment of injection.</li>
	<li>Before injection, transfer the aliquot to wet ice and keep it on ice at all times except for sonication and retrieval. The number and types of samples will be determined by the experimental design.</li>
	<li>Sonicate the selected samples 3x in a bath with floating wet ice to avoid warming up of the sample. Use 5 s cycles each time, waiting 1 min on wet ice in between cycles.</li>
	<li>Place 1 &#956;L of sample onto the thermoplastic. Make an indention in the surface, if needed, before sample placement to prevent movement of the sample.</li>
	<li>Use the knobs of the manipulator holding the nanoinjector to position the needle and move it toward the sample to be filled. Once the needle tip is in the sample, press and hold the FILL button until enough sample is taken up. About 0.5 &#956;L is required for 10 oocytes. Using the knobs, move the nanoinjector&#39;s needle back to the highest position.</li>
</ol>
3. Injecting the oocytes
NOTE: Standardized rat cortex synaptosomal membranes are also injected in all experiments into a set of oocytes to measure changes in fusion capacity between different batches of oocytes.
<ol>
	<li>Place selected oocytes onto the small Petri dish that has the fitted nylon mesh and Barth&#39;s solution (about 10 oocytes). Arrange oocytes so that they are not on top of each other; this will be helpful during the injection process.</li>
	<li>Using the knobs on the manipulator holding the nanoinjector, move the needle toward the oocytes. Once the needle is submerged in the Barth&#39;s solution, use the foot pedal for the nanoinjector to make sure that the microinjection needle is releasing sample. If the sample is being released it will be visible from the microscope view.</li>
	<li>Once it is confidently established that the sample is being released, move on to microinjecting the oocyte. If sample is not being released, repeat the process of filling the needle.</li>
	<li>Penetrate the oocyte just underneath the surface with the needle, no deeper, and use the foot pedal to inject the sample. The foot pedal will make a beep sound when the sample is released. Wait about 2-3 s. If the injection was successful, the cell will expand. Once this happens, use the knobs on the manipulator to exit the cell. 
	NOTE: Fusion of membranes in the oocyte is a polarized process; therefore, the oocyte should be injected in the animal side of the cell, preferably above the equator, with the angle of the needle toward the animal side to maximize membrane fusion.</li>
	<li>Move the Petri dish so that the next oocyte is aligned with the needle and repeat the process until all oocytes in the dish have been injected. Repeat this process until the desired number of oocytes has been injected. Make sure that one well of oocytes is left non-injected to be used as control.</li>
	<li>Once all oocytes have been injected, retract the nanoinjector to its original position and remove the needle.</li>
</ol>
4. Recording of ion currents using a two electrode voltage clamp
<ol>
	<li>To make two electrode voltage clamp (TEVC) electrode needles, pull 15 cm long fire polished borosilicate glass tubes by using the micropipette puller. Once pulling is complete, fill with 3 M KCl using long capillary needles, or alternatively, submerge and boil the needles in 3 M KCl solution for 15 min under a continuous vacuum. The high temperature helps in the filling of the electrodes and removal of air bubbles.
	<ol>
		<li>Prepare a 3 M KCl electrode fill solution by using KCl in the crystalline form and deionized water, and following these steps carefully so that reference potential of electrodes is not changed. Dry the KCl carefully in an oven for 2-3 h. Using an analytical balance, carefully weigh 223.68 g of KCl. Transfer the KCl to a 1 L glass bottle. Use this solution to chlorinate silver electrode wires. 
		NOTE: For pulling glass electrodes, the pipette cookbook can be downloaded here:&#160;https://www.sutter.com/PDFs/cookbook.pdf.</li>
	</ol>
	</li>
	<li>Turn on all equipment used for the recording of ion currents: main systems, solution valves, microscope light, valve systems, oocyte clamp, and vacuum system. Turn on the desktop computer, log into WinEDR V3.9.1., and make sure that the program is running and that the setup values are correct as follows: record to disk, set recording duration, and clear out the simulator option.
	<ol>
		<li>Provide the following initial setup values for the amplifier: for voltage electrode (Vm) section, turn off the negative capacity compensation (-C); for bath electrodes (Im) section, set the gain selector switch (range from 0.1 to 10) at 10, and the three-position toggle switch which selects the gain multiplier (x0.1, x1.0 and x10), at x1.0. The LED lights will indicate the gain multiplier selection; for clamp section, turn off the clamp mode selector, set the DC gain control to IN, and the gain control full-bandwidth open-loop (range 0 to 2000) to around 1200; for commands, set at 40mV and set the hold controls negative and scale multiplier to be in x2; for current electrode, use the Ve offset to establish a zero reference before impaling the oocyte.</li>
		<li>To record, open the WinEDR V3.9.1 software, go to the top menu, and select File &#62; Open New File. Create your own folder and save the file.</li>
		<li>Next, in the main menu select Record &#62; Record to Disk. When the two electrodes are inside the ooctyte, turn on the ringer in the VC-8 valve controller, and turn the clamp to Slow in the OC-725C amplifier. Then, go back to the WinEDR software and select Record at the top left of the screen.</li>
		<li>In the mark chart at the bottom left, write down your initial membrane potential and press Enter. As the experiment keeps going, you can write down the name of the drugs being used. Once the experiment is over, select Stop at the top left of the screen. 
		NOTE: We use WinEDR or WinWCP software to perform TEVC because these are free and well suited for experiments, but any other software that is available can be used.</li>
	</ol>
	</li>
	<li>Make up all of the drug solutions needed to perform the experiment (e.g., GABA, Kainate) and confirm that the solution valves are working properly by switching them on and off and observing the displacement of the pinch valves.</li>
	<li>Fill the tubes by pouring the corresponding solution on the syringe receptacle connected to the tube and label the valve controller to correlate with the solution tubes. Ensure that the flow of solution is steady, there are no bubbles and that the vacuum system is working properly.</li>
	<li>Set up the microscope and recording chamber area. Place the agar bridges (see below) in the respective circular holes behind the recording chamber (distal from handling area), connecting the wells for the electrodes used as ground reference and the recording chamber. Add Ringer&#39;s working solution (1x) to the recording chamber and the ground reference electrode wells.
	<ol>
		<li>Agar bridges are U-shaped borosilicate tubes, 2-3 cm long, filled with 3% agar in Ringer&#39;s solution. Make U-shaped tubes by cutting 15 cm long tubes, bending them on the open flame, and then filling them with hot 3% agar using an insulin syringe. Store filled agar bridges in Ringer&#39;s solution in the fridge until use.</li>
		<li>Make 20x Ringer&#39;s stock solution as follows: In 1 L of distilled water, add 134.4 g of NaCl, 2.982 g of KCl, 5.29 g of CaCl2,&#160;and 23.83 g of HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid). For 1x Ringer&#39;s working solution, in a volumetric flask, add 200 mL of Ringer&#39;s stock solution (20x) to 3,800 mL of distilled water.</li>
	</ol>
	</li>
	<li>Prepare chlorinated silver electrodes prior to each experiment by electrolysis by placing silver wires in 3 M KCl solution and connecting the silver electrode to the positive terminal of a 9 V battery. After about 3 min a solid brown layer of AgCl is deposited into the electrode. Place the chlorinated silver electrode wires into the electrode housing. 
	NOTE: Ground reference and recording chlorinated electrodes are silver/silver chloride (Ag/AgCl) electrodes.</li>
	<li>Remove the borosilicate needles from the KCl solution. Use a syringe to extract a small amount of KCl solution from the open end of the electrode needle and replace the KCl solution with mineral oil; this will avoid water evaporation and changes in KCl concentration within the needle. Slide the glass needle over the chlorinated silver electrode wire and tighten them in place.</li>
	<li>Using the right and left manipulators holding the electrodes, guide the electrode needles into the recording chamber that is filled with Ringer&#39;s Solution.</li>
	<li>Check the resistance of each electrode by zeroing out the Vm and Ve offset knobs and then pressing the Electrode Test buttons. The resistance should be between 0.5-3 M&#8486; (directly read as 5-30 mV in the screen). If the resistance is out of the range, replace using new microelectrodes.</li>
	<li>Fill the recording chamber with fresh Ringer&#39;s solution. Using a glass pipette, place a non-injected or a microtransplanted oocyte in the center of the recording chamber. Ensure that the oocyte is clearly visible under the microscope, with animal side up (see NOTE in step 3.4).</li>
	<li>Guide the electrode into the Ringer&#39;s solution until they touch the oocyte membrane. Gently pierce the oocyte membrane in the animal side (see NOTE in step 3.4) with both electrodes and record the resting membrane potential. Turn on the Ringer&#39;s solution flow.</li>
	<li>Using the oocyte clamp amplifier, change the mode to Voltage Clamp and set the holding voltage of -80 mV. The current on the monitor should be negative, usually between 0 and 0.4 microamperes.</li>
	<li>Create a new file to save the recording as File &#62; New &#62; Save the Recording on the software. Start recording, press Record &#62; Record to Disk. Add relevant information to the file by using the mark box dialog (test name, drugs used, etc.).</li>
	<li>Apply agonists (e.g., GABA, glutamate, or kainate) for chemical stimulation by opening the valves and perfusing them into the recording chamber for 15 s. Usually, use a gain of 10x to plot the maximum number of points and reconstruct the response. If currents are very small, increase the amplification, or gain, in order for them to be evaluated and visible. Always make note of these changes. 
	NOTE: The duration of the perfusion depends on the experimental paradigm. If the major goal is to measure maximum amplitude, 15 s will be enough to reach the peak for GABA or the plateau for kainate.</li>
	<li>Always monitor solution levels. Ringer&#39;s solution will need to be refilled frequently as it is used between all solution uses. Once recording is completed, turn the voltage clamp to the OFF position and turn the Ringer&#39;s solution valve off. Remove the oocyte. Stop recording and save the file. Repeat these steps for all the injected oocytes.</li>
</ol>
5. Analyzing TEVC recordings
<ol>
	<li>Save a copy of the recordings to a USB drive. From there, open the desired file to be analyzed. In the file that opens, the recording can be viewed, marked, and measured. Most common analysis includes measuring of maximum amplitude, activation time, desensitization, and rundown of repetitive applications.</li>
	<li>Measure the AMPA maximum response and the GABA peak response. To acquire these measurements, first establish a zero level or baseline by finding the red line with the cursor and dragging it to the current generated by Ringer application or baseline.</li>
	<li>Once the zero level is established, determine the response measurements by using the mouse to drag the green, vertical readout cursor to the desired part of the graph tracer. If wanted, export the traces to be analyzed in any other preferred software 
	NOTE: If the zero level is unable to be defined or there is too much noise indicated on the tracer, export the WinEDR file to an offline analytical software, that will allow for modifications to provide a level baseline and the addition of filters in order to reduce noise.</li>
</ol>

The authors have no conflicts of interest to disclose.

Analysis of native protein complexes from human brains is needed to understand homeostatic and pathological processes in brain disorders and develop therapeutic strategies to prevent or treat diseases. Thus, brain banks containing snap frozen samples are an invaluable source of a large and mostly untapped wealth of physiological information29,30. An initial concern to use postmortem tissue is the clear possibility of mRNA or protein degradation that may confound ...

Neurodegenerative disorders affect a large percentage of the population. Although their devastating consequences are well known, the link between the functional alterations of neurotransmitter receptors, which are critical for brain function, and their symptomatology is still poorly understood. Inter-individual variability, chronic nature of the disease, and insidious onset of symptoms are just some of the reasons that have delayed the understanding of the many brain disorders where chemical imbalances are well documented1,2. Animal models have generated invaluable information and expanded our knowledge about the mechanisms underlying physiology and pathophysiology in evolutionary conserved systems; however, several interspecies differences between rodents and humans preclude the direct extrapolation of receptor function from animal models to the human brain3. Thus, initial efforts to study native human receptors were developed by Ricardo Miledi&#39;s lab using surgically removed tissue and frozen samples. These initial experiments used whole membranes that include neuronal synaptic and extra synaptic receptors as well as non-neuronal neurotransmitter receptors, and although they provide important information about diseased states, there is a concern that the mix of receptors complicates the interpretation of data4,5,6,7. Importantly, synapses are the major target in many neurodegenerative disorders8,9; therefore, assays to test the functional properties of affected synapses are fundamental to obtain information about disease-relevant changes affecting synaptic communication. Here, a modification of the original method is described: microtransplantation of synaptic membranes (MSM), which focuses on the physiological characterization of enriched synaptic protein preparations and has been successfully applied to study rat and human synaptosomes10,11,12,13,14,15. With this methodology, it is possible to transplant synaptic receptors that were once working in the human brain, embedded in their own native lipids and with their own cohort of associated proteins. Moreover, because MSM data is quantitative, it is possible to use this data to integrate with large proteomic or sequencing datasets10.
It is important to note that many pharmacological and biophysical analyses of synaptic receptors are done on recombinant proteins16,17. While this approach provides better insight into the structure-function relationships of receptors, it cannot provide information about complex multimeric receptor complexes found in neurons and their changes in disease. Therefore, a combination of native and recombinant proteins should provide a more comprehensive analysis of synaptic receptors.
There are many methods to prepare synaptosomes10,11,12,13,14,15 which can be adjusted for the requirements of a lab. The protocol begins with the assumption that synaptosomal enriched preparations were isolated and are ready to be processed for microtransplantation experiments. In the lab, the Syn-Per method is used following the manufacturer instructions. This is done because of high reproducibility in electrophysiological experiments10,11. There is also abundant literature explaining how to isolate Xenopus oocytes18,19, which can also be purchased ready for injection20.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>For Microinjection</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>3.5&#34; Glass Capillaries</td>
			<td>Drummond</td>
			<td>3-000-203-G/X</td>
			<td></td>
		</tr>
		<tr>
			<td>24 well, flat bottom Tissue Culture Plate</td>
			<td>Thermofisher</td>
			<td>FB012929</td>
			<td></td>
		</tr>
		<tr>
			<td>Flaming/Brown type micropipette puller</td>
			<td>Sutter</td>
			<td>P-1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Injection Dish</td>
			<td>Thermofisher</td>
			<td>08-772B</td>
			<td></td>
		</tr>
		<tr>
			<td>Microcentrifuge Tubes</td>
			<td>Thermofisher</td>
			<td>02-682-002</td>
			<td></td>
		</tr>
		<tr>
			<td>Mineral Oil</td>
			<td>Thermofisher</td>
			<td>O121-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Nanoject II</td>
			<td>Drummond</td>
			<td>3-000-204</td>
			<td></td>
		</tr>
		<tr>
			<td>Nylon mesh</td>
			<td>Industrial Netting</td>
			<td>WN0800</td>
			<td></td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td>Thermofisher</td>
			<td>S37440</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereoscope</td>
			<td>Fisher Scientific</td>
			<td>03-000-037</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe</td>
			<td>Thermofisher</td>
			<td>14-841-31</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrasonic cleaning bath</td>
			<td>Thermofisher</td>
			<td>FS20D</td>
			<td></td>
		</tr>
		<tr>
			<td>Xenopus laevis frogs</td>
			<td>Xenopus 1</td>
			<td>4217</td>
			<td></td>
		</tr>
		<tr>
			<td>For Two Electrode Voltage clamp</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>15 cm long fire polished borosilicate glass capillaries</td>
			<td>Sutter</td>
			<td>B200-116-15</td>
			<td></td>
		</tr>
		<tr>
			<td>Any PC computer or laptop</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Low-pass Bessel Filter</td>
			<td>Warner Instruments</td>
			<td>LPF-8</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereoscope</td>
			<td>Fisher Scientific</td>
			<td>03-000-037</td>
			<td></td>
		</tr>
		<tr>
			<td>Two electrode voltage clamp workstation</td>
			<td>Warner Instruments</td>
			<td>TEV-700</td>
			<td></td>
		</tr>
		<tr>
			<td>ValveLink 8.2 Perfusion Controller</td>
			<td>Automate Scientific</td>
			<td>SKU:01-18</td>
			<td></td>
		</tr>
		<tr>
			<td>WInEDR Free software</td>
			<td>University of Strathclyde Glasgow</td>
			<td>https://spider.science.strath.ac.uk/sipbs/software_ses.htm</td>
			<td></td>
		</tr>
		<tr>
			<td>X Series Multifunction DAQ</td>
			<td>National Instruments</td>
			<td>NI USB-6341</td>
			<td></td>
		</tr>
		<tr>
			<td>Reagents</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium dichloride</td>
			<td>Thermofisher</td>
			<td>C79</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium nitrate tetrahydrate</td>
			<td>Thermofisher</td>
			<td>C109</td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase</td>
			<td>Sigma-Aldrich</td>
			<td>C0130</td>
			<td></td>
		</tr>
		<tr>
			<td>GABA</td>
			<td>Sigma-Aldrich</td>
			<td>A2129</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid)</td>
			<td>Thermofisher</td>
			<td>BP310</td>
			<td></td>
		</tr>
		<tr>
			<td>Kainic acid</td>
			<td>Tocris</td>
			<td>0222</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium sulfate heptahydrate</td>
			<td>Thermofisher</td>
			<td>M63</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium chloride</td>
			<td>Thermofisher</td>
			<td>P217</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium bicarbonate</td>
			<td>Thermofisher</td>
			<td>S233</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>Thermofisher</td>
			<td>S271-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrafree-0.1 &#181;m MC filter,</td>
			<td>Amicon</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

microtransplantation of synaptic membranes to reactivate human synaptic receptors for functional studies

Excitatory and inhibitory ionotropic receptors are the major gates of ion fluxes that determine the activity of synapses during physiological neuronal communication. Therefore, alterations in their abundance, function, and relationships with other synaptic elements have been observed as a major correlate of alterations in brain function and cognitive impairment in neurodegenerative diseases and mental disorders. Understanding how the function of excitatory and inhibitory synaptic receptors is altered by disease is of critical importance for the development of effective therapies. To gain disease-relevant information, it is important to record the electrical activity of neurotransmitter receptors that remain functional in the diseased human brain. So far this is the closest approach to assess pathological alterations in receptors' function. In this work, a methodology is presented to perform microtransplantation of synaptic membranes, which consists of reactivating synaptic membranes from snap frozen human brain tissue containing human receptors, by its injection and posterior fusion into the membrane of Xenopus laevis oocytes. The protocol also provides the methodological strategy to obtain consistent and reliable responses of &#945;-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) and &#947;-aminobutyric acid (GABA) receptors, as well as novel detailed methods that are used for normalization and rigorous data analysis.

Within a few hours after injection, the synaptic membranes, carrying their neurotransmitter receptors and ion channels, begin to fuse with the oocyte plasma membrane. Figure 1 shows recordings of AMPA and GABAA receptors microtransplanted into Xenopus oocytes. For most of the analysis, the responses from two or three oocytes per sample were measured, using two or three batches of oocytes from different frogs, for a total of six to nine oocytes per sample. This is done for...

Watch this Scientific Journal Video about Microtransplantation of Synaptic Membranes to Reactivate Human Synaptic Receptors for Functional Studies at JoVE.com

Microtransplantation of Synaptic Membranes to Reactivate Human Synaptic Receptors for Functional Studies

The protocol demonstrates that by performing microtransplantation of synaptic membranes into Xenopus laevis oocytes, it is possible to record consistent and reliable responses of &#945;-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid and &#947;-aminobutyric acid receptors.

Excitatory and inhibitory ionotropic receptors are the major gates of ion fluxes that determine the activity of synapses during physiological neuronal ...

microtransplantation-synaptic-membranes-to-reactivate-human-synaptic

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Neuroscience

2.1K Views.  University of Texas Medical Branch. The protocol demonstrates that by performing microtransplantation of synaptic membranes into Xenopus laevis oocytes, it is possible to record consistent and reliable responses of α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid and γ-aminobutyric acid receptors. 

Video: Microtransplantation of Synaptic Membranes to Reactivate Human Synaptic Receptors for Functional Studies

Studying Synaptic Vesicle Pools using Photoconversion of Styryl Dyes

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The vesicles are then retrieved from the plasma membrane (endocytosis) and grouped together with the general pool of vesicles within the nerve terminal, until they undergo a new exo- and endocytosis cycle (vesicle recycling). These processes have been studied using a variety of techniques such as electron microscopy, electrophysiology recordings, amperometry and capacitance measurements. Importantly, during the last two decades a number of fluorescently labeled markers emerged, allowing optical techniques to track vesicles in their recycling dynamics. One of the most commonly used markers is the styryl or FM dye 1; structurally, all FM dyes contain a hydrophilic head and a lipophilic tail connected through an aromatic ring and one or more double bonds (Fig. 1B). A classical FM dye experiment to label a pool of vesicles consists in bathing the preparation (Fig. 1Ai) with the dye during the stimulation of the nerve (electrically or with high K+). This induces vesicle recycling and the subsequent loading of the dye into recently endocytosed vesicles (Fig. 1Ai-iii). After loading the vesicles with dye, a second round of stimulation in a dye-free bath would trigger the FM release through exocytosis (Fig. 1Aiv-v), process that can be followed by monitoring the fluorescence intensity decrease (destaining).

Although FM dyes have contributed greatly to the field of vesicle recycling, it is not possible to determine the exact localization or morphology of individual vesicles by using conventional fluorescence microscopy. For that reason, we explain here how FM dyes can also be used as endocytic markers using electron microscopy, through photoconversion. The photoconversion technique exploits the property of fluorescent dyes to generate reactive oxygen species under intense illumination. Fluorescently labeled preparations are submerged in a solution containing diaminobenzidine (DAB) and illuminated. Reactive species generated by the dye molecules oxidize the DAB, which forms a stable, insoluble precipitate that has a dark appearance and can be easily distinguished in electron microscopy 2,3. As DAB is only oxidized in the immediate vicinity of fluorescent molecules (as the reactive oxygen species are short-lived), the technique ensures that only fluorescently labeled structures are going to contain the electron-dense precipitate. The technique thus allows the study of the exact location and morphology of actively recycling organelles.

FM dyes have been of invaluable help in the understanding of synaptic dynamics. FMs are normally followed under the fluorescent microscope during different stimulation conditions. However, photoconversion of FM dyes combined with electron microscopy allows the visualization of distinct synaptic vesicle pools, among other ultrastructure components, in synaptic boutons.

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The ...

Lateral Diffusion and Exocytosis of Membrane Proteins in Cultured Neurons Assessed using Fluorescence Recovery and Fluorescence-loss Photobleaching

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This directed trafficking is of fundamental importance to deliver, maintain or remove synaptic proteins.
Super-ecliptic pHluorin (SEP) is a pH-sensitive derivative of eGFP that has been extensively used for live cell imaging of plasma membrane proteins1-2. At low pH, protonation of SEP decreases photon absorption and eliminates fluorescence emission. As most intracellular trafficking events occur in compartments with low pH, where SEP fluorescence is eclipsed, the fluorescence signal from SEP-tagged proteins is predominantly from the plasma membrane where the SEP is exposed to a neutral pH extracellular environment. When illuminated at high intensity SEP, like every fluorescent dye, is irreversibly photodamaged (photobleached)3-5. Importantly, because low pH quenches photon absorption, only surface expressed SEP can be photobleached whereas intracellular SEP is unaffected by the high intensity illumination6-10. 
FRAP (fluorescence recovery after photobleaching) of SEP-tagged proteins is a convenient and powerful technique for assessing protein dynamics at the plasma membrane. When fluorescently tagged proteins are photobleached in a region of interest (ROI) the recovery in fluorescence occurs due to the movement of unbleached SEP-tagged proteins into the bleached region. This can occur via lateral diffusion and/or from exocytosis of non-photobleached receptors supplied either by de novo synthesis or recycling (see Fig. 1). The fraction of immobile and mobile protein can be determined and the mobility and kinetics of the diffusible fraction can be interrogated under basal and stimulated conditions such as agonist application or neuronal activation stimuli such as NMDA or KCl application8,10. 
We describe photobleaching techniques designed to selectively visualize the recovery of fluorescence attributable to exocytosis. Briefly, an ROI is photobleached once as with standard FRAP protocols, followed, after a brief recovery, by repetitive bleaching of the flanking regions. This 'FRAP-FLIP' protocol, developed in our lab, has been used to characterize AMPA receptor trafficking at dendritic spines10, and is applicable to a wide range of trafficking studies to evaluate the intracellular trafficking and exocytosis.

This report describes the use of live cell imaging and photobleach techniques to determine the surface expression, transport pathways and trafficking kinetics of exogenously expressed, pH-sensitive GFP-tagged proteins at the plasma membrane of neurons.

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This ...

Examination of Synaptic Vesicle Recycling Using FM Dyes During Evoked, Spontaneous, and Miniature Synaptic Activities

Synaptic vesicles in functional nerve terminals undergo exocytosis and endocytosis. This synaptic vesicle recycling can be effectively analyzed using styryl FM dyes, which reveal membrane turnover. Conventional protocols for the use of FM dyes were designed for analyzing neurons following stimulated (evoked) synaptic activity. Recently, protocols have become available for analyzing the FM signals that accompany weaker synaptic activities, such as spontaneous or miniature synaptic events. Analysis of these small changes in FM signals requires that the imaging system is sufficiently sensitive to detect small changes in intensity, yet that artifactual changes of large amplitude are suppressed. Here we describe a protocol that can be applied to evoked, spontaneous, and miniature synaptic activities, and use cultured hippocampal neurons as an example. This protocol also incorporates a means of assessing the rate of photobleaching of FM dyes, as this is a significant source of artifacts when imaging small changes in intensity.

We describe the use of styryl FM dyes to image synaptic vesicle recycling in functional nerve terminals. This protocol can be applied not only to evoked, but also spontaneous and miniature synaptic activities. The protocol expands the variety of synaptic events that can be effectively evaluated.

Synaptic vesicles in functional nerve terminals undergo exocytosis and endocytosis. This synaptic vesicle recycling can be effectively analyzed using ...

Deriving the Time Course of Glutamate Clearance with a Deconvolution Analysis of Astrocytic Transporter Currents

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the membrane with the co-transport of 3 Na+ and 1 H+ and the counter-transport of 1 K+. The stoichiometric current generated by the transport process can be monitored with whole-cell patch-clamp recordings from astrocytes. The time course of the recorded current is shaped by the time course of the glutamate concentration profile to which astrocytes are exposed, the kinetics of glutamate transporters, and the passive electrotonic properties of astrocytic membranes. Here we describe the experimental and analytical methods that can be used to record glutamate transporter currents in astrocytes and isolate the time course of glutamate clearance from all other factors that shape the waveform of astrocytic transporter currents. The methods described here can be used to estimate the lifetime of flash-uncaged and synaptically-released glutamate at astrocytic membranes in any region of the central nervous system during health and disease.

We describe an analytical method to estimate the lifetime of glutamate at astrocytic membranes from electrophysiological recordings of glutamate transporter currents in astrocytes.

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the ...

High-Resolution Quantitative Immunogold Analysis of Membrane Receptors at Retinal Ribbon Synapses

Retinal ganglion cells (RGCs) receive excitatory glutamatergic input from bipolar cells. Synaptic excitation of RGCs is mediated postsynaptically by NMDA receptors (NMDARs) and AMPA receptors (AMPARs). Physiological data have indicated that glutamate receptors at RGCs are expressed not only in postsynaptic but also in perisynaptic or extrasynaptic membrane compartments. However, precise anatomical locations for glutamate receptors at RGC synapses have not been determined. Although a high-resolution quantitative analysis of glutamate receptors at central synapses is widely employed, this approach has had only limited success in the retina. We developed a postembedding immunogold method for analysis of membrane receptors, making it possible to estimate the number, density and variability of these receptors at retinal ribbon synapses. Here we describe the tools, reagents, and the practical steps that are needed for: 1) successful preparation of retinal fixation, 2) freeze-substitution, 3) postembedding immunogold electron microscope (EM) immunocytochemistry and, 4) quantitative visualization of glutamate receptors at ribbon synapses.

The postembedding immunogold method is one of the most effective ways to provide high-resolution analyses of the subcellular localization of specific molecules. Here we describe a protocol to quantitatively analyze glutamate receptors at retinal ribbon synapses.

Retinal ganglion cells (RGCs) receive excitatory glutamatergic input from bipolar cells. Synaptic excitation of RGCs is mediated postsynaptically by NMDA ...

Analyzing Synaptic Modulation of Drosophila melanogaster Photoreceptors after Exposure to Prolonged Light

The nervous system has the remarkable ability to adapt and respond to various stimuli. This neural adjustment is largely achieved through plasticity at the synaptic level. The Active Zone (AZ) is the region at the presynaptic membrane that mediates neurotransmitter release and is composed of a dense collection of scaffold proteins. AZs of Drosophila melanogaster (Drosophila) photoreceptors undergo molecular remodeling after prolonged exposure to natural ambient light. Thus the level of neuronal activity can rearrange the molecular composition of the AZ and contribute to the regulation of the functional output.
Starting from the light exposure set-up preparation to the immunohistochemistry, this protocol details how to quantify the number, the spatial distribution, and the delocalization level of synaptic molecules at AZs in Drosophila photoreceptors. Using image analysis software, clusters of the GFP-fused AZ component Bruchpilot were identified for each R8 photoreceptor (R8) axon terminal. Detected Bruchpilot spots were automatically assigned to individual R8 axons. To calculate the distribution of spot frequency along the axon, we implemented a customized software plugin. Each axon&#39;s start-point and end-point were manually defined and the position of each Bruchpilot spot was projected onto the connecting line between start and end-point. Besides the number of Bruchpilot clusters, we also quantified the delocalization level of Bruchpilot-GFP within the clusters. These measurements reflect in detail the spatially resolved synaptic dynamics in a single neuron under different environmental conditions to stimuli.

Analyzing Synaptic Modulation of Drosophila melanogaster Photoreceptors after Exposure to Prolonged Light

Here we show how to quantify the number and spatial distribution of synaptic active zones in Drosophila melanogaster&#160;photoreceptors, highlighted with a genetically encoded molecular marker, and their modulation after prolonged exposure to light.

The nervous system has the remarkable ability to adapt and respond to various stimuli. This neural adjustment is largely achieved through plasticity at ...

Culturing In Vivo-like Murine Astrocytes Using the Fast, Simple, and Inexpensive AWESAM Protocol

The AWESAM (a low-cost easy stellate astrocyte method) protocol entails a fast, simple, and inexpensive way to generate large quantities of in vivo-like mouse and rat astrocyte monocultures: Brain cells can be isolated from different brain regions, and after a week of cell culture, non-astrocytic cells are shaken off by placing the culture dishes on a shaker for 6 h in the incubator. The remaining astrocytes are then passaged into new plates with an astrocyte-specific medium (termed NB+H). NB+H contains low concentrations of heparin-binding EGF-like growth factor (HBEGF), which is used in place of serum in medium. After growing in NB+H, AWESAM astrocytes have a stellate morphology and feature fine processes. Moreover, these astrocytes have more in vivo-like gene expression than astrocytes generated by previously published methods. Ca2+ imaging, vesicle dynamics, and other events close to the membrane can thus be studied in the fine astrocytic processes in vitro, e.g., using live cell confocal or TIRF microscopy. Notably, AWESAM astrocytes also exhibit spontaneous Ca2+ signaling similar to astrocytes in vivo.

Culturing In Vivo-like Murine Astrocytes Using the Fast, Simple, and Inexpensive AWESAM Protocol

The AWESAM protocol described here is optimal for culturing murine astrocytes in isolation from other brain cells in a fast, simple, and inexpensive manner. AWESAM astrocytes exhibit spontaneous Ca2+ signaling, morphology, and gene expression profiles similar to astrocytes in vivo.

The AWESAM (a low-cost easy stellate astrocyte method) protocol entails a fast, simple, and inexpensive way to generate large quantities of in vivo-like ...

Pentylenetetrazole-Induced Kindling Mouse Model

Pentylenetetrazole (PTZ) is a GABA-A receptor antagonist. An intraperitoneal injection of PTZ into an animal induces an acute, severe seizure at a high dose, whereas sequential injections of a subconvulsive dose have been used for the development of chemical kindling, an epilepsy model. A single low-dose injection of PTZ induces a mild seizure without convulsion. However, repetitive low-dose injections of PTZ decrease the threshold to evoke a convulsive seizure. Finally, continuous low-dose administration of PTZ induces a severe tonic-clonic seizure. This method is simple and widely applicable to investigate the pathophysiology of epilepsy, which is defined as a chronic disease that involves repetitive seizures. This chemical kindling protocol causes repetitive seizures in animals. With this method, vulnerability to PTZ-mediated seizures or the degree of aggravation of epileptic seizures was estimated. These advantages have led to the use of this method for screening anti-epileptic drugs and epilepsy-related genes. In addition, this method has been used to investigate neuronal damage after epileptic seizures because the histological changes observed in the brains of epileptic patients also appear in the brains of chemical-kindled animals. Thus, this protocol is useful for conveniently producing animal models of epilepsy.

This protocol describes a method of chemical kindling with pentylenetetrazole and provides a mouse model of epilepsy. This protocol can also be used to investigate vulnerability to seizure induction and pathogenesis after epileptic seizures in mice.

Pentylenetetrazole (PTZ) is a GABA-A receptor antagonist. An intraperitoneal injection of PTZ into an animal induces an acute, severe seizure at a high ...

Combining Optogenetics with Artificial microRNAs to Characterize the Effects of Gene Knockdown on Presynaptic Function within Intact Neuronal Circuits

The purpose of this protocol is to characterize the effect of gene knockdown on presynaptic function within intact neuronal circuits. We describe a workflow on how to combine artificial microRNA (miR)-mediated RNA interference with optogenetics to achieve selective stimulation of manipulated presynaptic boutons in acute brain slices. The experimental approach involves the use of a single viral construct and a single neuron-specific promoter to drive the expression of both an optogenetic probe and artificial miR(s) against presynaptic gene(s). When stereotactically injected in the brain region of interest, the expressed construct makes it possible to stimulate with light exclusively the neurons with reduced expression of the gene(s) under investigation. This strategy does not require the development and maintenance of genetically modified mouse lines and can in principle be applied to other organisms and to any neuronal gene of choice. We have recently applied it to investigate how the knockdown of alternative splice isoforms of presynaptic P/Q-type voltage-gated calcium channels (VGCCs) regulates short-term synaptic plasticity at CA3 to CA1 excitatory synapses in acute hippocampal slices. A similar approach could also be used to manipulate and probe the neuronal circuitry in vivo.

This protocol provides a workflow on how to combine artificial microRNA-mediated RNA interference with optogenetics to stimulate specifically presynaptic boutons with reduced expression of selective gene(s) within intact neuronal circuits.

The purpose of this protocol is to characterize the effect of gene knockdown on presynaptic function within intact neuronal circuits. We describe a ...

Single Synapse Indicators of Glutamate Release and Uptake in Acute Brain Slices from Normal and Huntington Mice

Synapses are highly compartmentalized functional units that operate independently on each other. In Huntington&#39;s disease (HD) and other neurodegenerative disorders, this independence might be compromised due to insufficient glutamate clearance and the resulting spill-in and spill-out effects. Altered astrocytic coverage of the presynaptic terminals and/or dendritic spines as well as a reduced size of glutamate transporter clusters at glutamate release sites have been implicated in the pathogenesis of diseases resulting in symptoms of dys-/hyperkinesia. However, the mechanisms leading to the dysfunction of glutamatergic synapses in HD are not well understood. Improving and applying synapse imaging we have obtained data shedding new light on the mechanisms impeding the initiation of movements. Here, we describe the principle elements of a relatively inexpensive approach to achieve single synapse resolution by using the new genetically encoded ultrafast glutamate sensor iGluu, wide-field optics, a scientific CMOS (sCMOS) camera, a 473 nm laser and a laser positioning system to evaluate the state of corticostriatal synapses in acute slices from age appropriate healthy or diseased mice. Glutamate transients were constructed from single or multiple pixels to obtain estimates of i) glutamate release based on the maximal elevation of the glutamate concentration [Glu] next to the active zone and ii) glutamate uptake as reflected in the time constant of decay (TauD) of the perisynaptic [Glu]. Differences in the resting bouton size and contrasting patterns of short-term plasticity served as criteria for the identification of corticostriatal terminals as belonging to the intratelencephalic (IT) or the pyramidal tract (PT) pathway. Using these methods, we discovered that in symptomatic HD mice ~40% of PT-type corticostriatal synapses exhibited insufficient glutamate clearance, suggesting that these synapses might be at risk to excitotoxic damage. The results underline the usefulness of TauD as a biomarker of dysfunctional synapses in Huntington mice with a hypokinetic phenotype.

We present a protocol to evaluate the balance between glutamate release and clearance at single corticostriatal glutamatergic synapses in acute slices from adult mice. This protocol uses the fluorescent sensor iGluu for glutamate detection, a sCMOS camera for signal acquisition and a device for focal laser illumination.