This work was supported by Ministerio de Econom&#236;a y Competitividad/Instituto de Salud Carlos III (SAF2014-55700-P, PCIN-2013-019-C03-03 and PIE14/00034), Instituci&#242; Catalana de Recerca i Estudis Avan&#231;ats (ICREA Academia-2010), and Agentschap voor Innovatie door Wetenschap en Technologie (SBO-140028) to F. C. Also, X. M, V. F.-D., and F. C. belong to the "Neuropharmacology and Pain" accredited research group (Generalitat de Catalunya,2014 SGR 1251). The work was also supported by the "Programa Pesquisador Visitante Especial-Ci&#234;ncia sem Fronteiras" from CAPES (Brazil) to F.C.

All animal experimental procedures were approved by the University of Barcelona Committee on Animal Use and Care (CEEA), in compliance with the guidelines described in the Guide for the Care and Use of Laboratory Animals11 and following the European Community, law 86/609/CCE, FELASA, and ARRIVE guidelines. Thus, mice are housed in standard cages, with ad libitum access to food and water, and are maintained under controlled standard conditions (12 h dark/light cycle starting at 7:30 AM, 22 &#176;C temperature, and 66% humidity).
1. Obtaining Mouse Brain Synaptosomes Using a Discontinuous Sucrose Gradient
Note: This method was reported previously10.
<ol>
	<li>Prepare fresh isolation buffer (IB), pH 7.4; 2 M sucrose; 1 M sucrose/0.1 mM CaCl2; and 0.1 mM CaCl2 (see Table 1). Chill the solutions on ice.</li>
	<li>Sacrifice five mice by cervical dislocation. Decapitate and rapidly remove the brain of each animal. Dissect the brain region of interest (i.e., the hippocampus, 0.25-0.35 g of fresh tissue) and place it in a 5 mL Potter-Elvehjem glass tube on ice with 1 mL of IB.</li>
	<li>Homogenize the tissue using a homogenizing stirrer (10 strokes at 700-900 rotations per min), preferentially in an ice bath to preclude sample warming.</li>
	<li>Place the homogenized tissue (1 mL) into a 15 mL tube containing 6 mL of 2 M sucrose and 2.5 mL of 0.1 mM CaCl2 at 4 &#176;C. Mix slowly by inverting the tube. 
	Note: The addition of calcium is essential to maintaining the adhesive interactions between organelles and thus to preserving the structure of the different &#34;densities.&#34;</li>
	<li>Place the solution in a 12 mL centrifuge tube and slowly add 2.5 mL of 1 M sucrose/0.1 mM CaCl2 on top of each tube to form the sucrose gradient. 
	Note: Label the position of the gradient interface on the centrifuge tube using a permanent marker.</li>
	<li>Weigh and equilibrate the centrifuge tubes with IB solution within their corresponding steel supports and with their closing lids.</li>
	<li>Centrifuge the tubes for 3 h at 100,000 x g and 4 &#176;C using a swinging bucket rotor centrifuge. Completely fill the rotor with all the steel supports (even empty, if required).</li>
	<li>Discard the top layer containing myelin. With a Pasteur pipette, collect the white ring between the 1.25-M and 1-M sucrose interphase corresponding to the synaptosome fraction.</li>
	<li>Dilute the synaptosomes with 9X their volume using IB in a centrifuge tube.</li>
	<li>Weigh and equilibrate the centrifuge tubes with IB solution within their corresponding steel supports and with their closing lids.</li>
	<li>Centrifuge the tubes for 30 min at 15,000 x g and 4 &#176;C using a swinging bucket rotor centrifuge.</li>
	<li>Discard the supernatant and resuspend the pellet with 1.1 mL of IB solution. Collect 100 &#181;L of this synaptosomal solution and centrifuge for 5 min at 11,000 x g and 4 &#186;C.</li>
	<li>Resuspend the synaptosomal pellet in 5% sodium dodecyl sulfate (SDS) and store this sample at -20 &#186;C. 
	Note: This sample will correspond to the total synaptosome fraction for further Western-blot analysis. The remaining synaptosomal fraction (~1 mL) can either be frozen down at -20 &#186;C until use or processed for subsequent subsynaptic fractionation. The synaptosomes or purified synapses represent between 1 and 2% of the total hippocampal volume12.</li>
</ol>
2. Pre-, Post- and Extrasynaptic-isolation
<ol>
	<li>Prepare 2x fresh solubilization buffer, pH 6.0; 1x solubilization buffer, pH 6.0; solubilization buffer, pH 8.0; and 0.1 mM CaCl2 (see Table 1). Chill the solutions on ice. 
	Note: The pH of these solutions should be accurately adjusted to achieve a good subsynaptic fractionation.</li>
	<li>Slowly dilute the 1-mL resuspended synaptosomal fraction (see step 1.12) with 5 mL of 0.1 mM CaCl2.</li>
	<li>Add the same volume (5 mL) of ice-cold 2x solubilization buffer, pH 6.0, and incubate for 50 min on ice under high agitation in a beaker on ice. 
	Note: While 1% of Triton X-100 at pH 6.0 solubilizes all plasma membrane proteins, it preserves proteins within the synaptic contacts or junctions (i.e., pre- and postsynaptic structures)4.</li>
	<li>Place the solution in a centrifuge tube. Weigh and equilibrate the tubes with equivalent volumes of 0.1 mM CaCl2 and 2x solubilization buffer solution, pH 6.0, within their corresponding steel supports and with their closing lids.</li>
	<li>Centrifuge the tubes for 30 min at 40,000 x g and 4 &#176;C using a swinging bucket rotor centrifuge. The resulting supernatant represents the extrasynaptic fraction, and the pellet corresponds to the synaptic junctions (i.e., pre- and postsynaptic fractions).</li>
	<li>Concentrate the supernatant containing the extrasynaptic fraction to a final 200 &#181;L volume using a 15 mL 10K filter tube centrifuged at 4,000 x g and 4 &#186;C.</li>
	<li>Precipitate the resulting concentrated extrasynaptic fraction (200 &#181;L) with 5 volumes (1 mL) of pre-chilled (-20 &#176;C) acetone overnight at -20 &#176;C.</li>
	<li>The next day, centrifuge the extrasynaptic fraction for 30 min at 18,000 x g and -15 &#176;C. After centrifugation, discard the acetone supernatant and dry the pellet to remove any trace of acetone.</li>
	<li>Finally, resuspend the pellet containing the extrasynaptic proteins with 200 &#181;L of 5% SDS. Sonicate the pellet, if required.</li>
	<li>Without disrupting it, carefully wash the pellet containing the pre- and postsynaptic fractions with 2 mL of 1x solubilization buffer, pH 6.0. Discard the buffer.</li>
	<li>Resuspend the pellet with 10 mL of 1x solubilization buffer, pH 8.0, using a glass Pasteur pipette. 
	Note: While 1% of Triton X-100 at pH 8.0 solubilizes the presynaptic specialization, it preserves the insoluble postsynaptic density4.</li>
	<li>Incubate the suspension for 50 min on ice under high agitation in a beaker on ice.</li>
	<li>Place the solution in a centrifuge tube. Weigh and equilibrate the tubes with 1x solubilization buffer solution, pH 8.0, within their corresponding steel supports and with their closing lids.</li>
	<li>Centrifuge the tubes for 30 min at 40,000 x g and 4 &#176;C using a swinging bucket rotor centrifuge; the resulting supernatant corresponds to the presynaptic fraction and the pellet to the postsynaptic fraction.</li>
	<li>Process the supernatant containing the presynaptic fraction, as described in steps 2.6-2.9.</li>
	<li>Resuspend the pellet containing postsynaptic fraction with 200 &#181;L of 5% SDS. Sonicate the pellet, if required.</li>
</ol>
3. Analyze the samples by immunoblot to validate the membrane fractionation
<ol>
	<li>Determine the amount of protein in each fraction (i.e., the total, extra-, pre- and post-synaptic fractions) using the bicinchoninic acid protein assay.</li>
	<li>Take 20 &#181;g of protein from each fraction and dilute it to a final volume of 50 &#181;L in SDS-polyacrylamide gel electrophoresis (SDS-PAGE) sample buffer. Boil for 5 min at 100 &#186;C.</li>
	<li>Separate the proteins by SDS-PAGE electrophoresis 10%, with a 4% concentrating gel under reducing conditions. Electrophorese at constant voltage of 80 V until the dye enters the lower gel and then increase the voltage to 120 V.</li>
	<li>Transfer the proteins to a PVDF membrane and block with IB blocking solution for 45 min at room temperature under continuous shaking.</li>
	<li>Incubate the membrane overnight at 4 &#186;C with the indicated primary antibody (i.e., anti-SNAP-25, anti-PSD-95, anti-synaptophysin, or anti-GPR37) diluted in IB blocking solution under continuous shaking.</li>
	<li>Wash the membrane three times (10 min each) with IB washing solution to eliminate unbound primary antibody.</li>
	<li>Incubate with the indicated secondary antibody for 90 min in dark conditions at room temperature under continuous shaking.</li>
	<li>Wash the membrane three times (10 min each) with IB washing solution to eliminate unbound secondary antibody.</li>
	<li>Incubate the membrane with chemiluminescent substrate (prepare the mix under dark conditions and following the 1:1 proportion of solution A and B provided by the manufacturer).</li>
	<li>Analyze the membrane in a chemiluminescent detection device.</li>
</ol>

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The authors have nothing to disclose.

The protocol presented here constitutes a powerful biochemical tool for the study of the subsynaptic distribution of specific proteins within any brain region. However, there are some drawbacks inherent to the technique that deserve to be highlighted here. For instance, one of the main limitations is the relatively large amount of tissue needed to purify a reasonable amount of protein in order to perform the immunoblot analysis of all subsynaptic fractions. This issue might be related to the fact that synapses (i.e.,...

Synaptic transmission relies on the physical integrity of the synapse, a concept that was envisaged as early as 1897 by Foster and Sherrington1. Thus, understanding the distribution of key neurotransmission components (e.g., ion channels, receptors, etc.) is essential to elucidate synaptic function, both in normal and pathological conditions. Electron microscopy (EM) has contributed enormously to the current ultrastructural notion of prototypical central nervous system (CNS) synapses. In such a way, EM has finely established the differences between pre- and postsynaptic densities, which are separated by a cleft of a rather uniform distance (~25 nm)2. Interestingly, the postsynaptic apparatus exhibits a relatively continuous, electron-dense thickening below its plasma membrane, the so-called postsynaptic density, or PSD2. Conversely, at the presynaptic apparatus, a noticeable discontinuous cytomatrix network is arranged just beneath the plasma membrane, which is essential to the alignment and docking of synaptic vesicles to the plasma membrane active zone3. Hence, EM constitutes the golden experimental approach to survey the distribution of proteins within structurally preserved CNS synapses. However, the information provided by electron micrographs is static. Indeed, accumulating evidences show that in vivo synapses are extremely dynamic, thus experiencing dramatic structural changes upon sustained synaptic transmission. In addition, the morphology and composition of synapses can change throughout different CNS regions and upon development, maturation, aging, and the development of neuropathological conditions. Overall, a protocol focused on isolating proteins belonging to different synaptic compartments in physiological conditions represents a valuable tool for a more comprehensive study of synaptic functioning.
Here, we describe this kind of complementary experimental approach, which allows for the preparative biochemical enrichment of the different synaptic membrane compartments-namely, extra-, pre- and postsynaptic membrane domains. This membrane fractionation method, first described by Philips et al. (2001)4, is based on a pH shift that weakens the adhesive interactions occurring within the pre- and postsynaptic apparatus. First, by using mild detergents at pH 6.0, it is possible to discern the adherens junction that holds the pre- and postsynaptic apparatus and that is maintained from the extrasynaptic membrane domain, which is solubilized and thus can be extracted from the synaptic contacts. Subsequently, raising the pH from 6.0 to 8.0 in the presence of mild detergents weakens the strength of the adherens junction that keeps the presynaptic active zone tightly bound to the postsynaptic density. Hence, the presynaptic compartment is solubilized and can be separated from the postsynaptic density, which is mostly preserved because the concentration of detergent used does not promote its solubilization4. Interestingly, the fractionation efficiency, eventually higher than 90%, can be confirmed by different subsynaptic markers: i) synaptosomal-associated protein 25 (SNAP-25), from the presynaptic active zone; ii) synaptophysin, from the extrasynaptic fraction (i.e., outside the active zone and including microsomes); and iii) postsynaptic density protein 95 (PSD-95), from the postsynaptic density. Notably, this brain membrane fractionation method has been used successfully. Accordingly, it has been possible to precisely determine the subsynaptic localization of different receptors, such as alpha-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) receptors5, adenosine A1 receptor (A1R)6, adenosine A2A receptor (A2AR)7, adenosine triphosphate (ATP) P2 receptors8, nicotinic acetylcholine receptor subunits9, and Parkinson&#39;s disease-associated receptor GPR3710. However, a number of limitations may impede the proper assessment of the synaptic distribution of a particular neuronal protein. Thus, in this procedure, we not only fully describe the entire protocol, but we also highlight some critical points to be considered, such as the rather large amount of tissue needed, the low protein yield, and the mandatory requirement to validate the efficiency of each separation before performing the definite experiment.

<table><tbody><tr><td>Sucrose</td><td>Pancreac Qu&amp;iacute;mica SL, Barcelona, Spain</td><td>1,316,211,211</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Pancreac Qu&amp;iacute;mica SL, Barcelona, Spain</td><td>2,112,211,210</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;6H2O</td><td>Pancreac Qu&amp;iacute;mica SL, Barcelona, Spain</td><td>1,313,961,210</td><td></td></tr><tr><td>Protease inhibitor cocktail Set III</td><td>Millipore, Darmstadt, Germany</td><td>535140</td><td></td></tr><tr><td>Trizma Base</td><td>Sigma, St. Louis, MO, USA</td><td>T1503</td><td></td></tr><tr><td>Tris-HCl</td><td>Pancreac Qu&amp;iacute;mica SL, Barcelona, Spain</td><td>1,236,541,209</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma, St. Louis, MO, USA</td><td>X100</td><td></td></tr><tr><td>SDS</td><td>Sigma, St. Louis, MO, USA</td><td>L3771</td><td></td></tr><tr><td>Glycerol</td><td>Sigma, St. Louis, MO, USA</td><td>G5516</td><td></td></tr><tr><td>Bromophenol Blue</td><td>Pancreac Qu&amp;iacute;mica SL, Barcelona, Spain</td><td>1,311,651,604</td><td></td></tr><tr><td>Dithiothreitol</td><td>Sigma, St. Louis, MO, USA</td><td>D0632</td><td></td></tr><tr><td>Tween 20</td><td>Sigma, St. Louis, MO, USA</td><td>P2287</td><td></td></tr><tr><td>Non fat dry milk</td><td></td><td></td><td></td></tr><tr><td>NaCl</td><td>Pancreac Qu&amp;iacute;mica SL, Barcelona, Spain</td><td>1,216,591,211</td><td></td></tr><tr><td>KCl</td><td>Pancreac Qu&amp;iacute;mica SL, Barcelona, Spain</td><td>1,314,941,210</td><td></td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Merck</td><td>4873</td><td></td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>Pancreac Qu&amp;iacute;mica SL, Barcelona, Spain</td><td>1,316,781,211</td><td></td></tr><tr><td>Basic 20 pH</td><td>Crison, Alella, Spain</td><td></td><td></td></tr><tr><td>Polytron VDI 12 Adaptable Homogenizer</td><td>VWR, Radnor, PA, USA.</td><td></td><td></td></tr><tr><td>Ultra-Clear Tubes (14x89mm)</td><td>Beckman Coulter, Hospitalet de Llobregat, Barcelona</td><td>344059</td><td>Tubes should be filled almost completely when used to prevent collapsing due to ultracentrifugation.</td></tr><tr><td>Amicon Ultra-15 Centrifugal filters Ultracel -10K</td><td>Merck Millipore, Darmstadt, Germany</td><td>UFC901008</td><td></td></tr><tr><td>Centrifuge 5430R</td><td>Eppendorf, Hamburgo, Germany</td><td></td><td></td></tr><tr><td>Optima L-90K Ultracentrifuge</td><td>Beckman Coulter, Hospitalet de Llobregat, Barcelona</td><td></td><td></td></tr><tr><td>Sonifier 250</td><td>Branson, Danbury, Connecticut</td><td></td><td></td></tr><tr><td>Amersham Imager 600</td><td>GE Healthcare Europe GmbH, Barcelona, Spain</td><td></td><td></td></tr><tr><td>Disposable Glass Pasteur Pippetes 230 mm</td><td>VWR, Radnor, PA, USA</td><td>612-1702</td><td></td></tr><tr><td>Compact Balance EK-610</td><td>A&amp;amp;D, Tokyo, Japan</td><td></td><td></td></tr><tr><td>Pierce&amp;trade; BCA Protein Assay Kit</td><td>Pierce Biotechnology, Rockford, IL, USA</td><td></td><td></td></tr><tr><td>SuperSignal west pico chemiluminescent substrate</td><td>Thermo Fisher Scientific Inc., Waltham, MA, USA</td><td></td><td></td></tr><tr><td>GR 200 Precision Balance</td><td>A&amp;amp;D, Tokyo, Japan</td><td></td><td></td></tr><tr><td>Anti-GPR37</td><td>Homemade antibody anti-GPR37 produced and validated in Francisco Ciruela Laboratory.</td><td></td><td>Primary antibodies used at a final concentration of 0.250 &amp;mu;g/mL</td></tr><tr><td>Anti-SNAP-25, anti-PSD-95, anti-synaptophysin</td><td>Abcam, Cambridge, United Kingdom</td><td></td><td>Primary antibodies diluted 1:10,000</td></tr><tr><td>HRP-conjugated goat anti-mouse IgG</td><td>Thermo Fisher Scientific, Waltham, MA, USA.</td><td></td><td>Secondary antibody diluted 1:10,000</td></tr><tr><td>HRP-conjugated goat anti-rabbit IgG</td><td>Thermo Fisher Scientific, Waltham, MA, USA.</td><td></td><td>Secondary antibody diluted 1:30,000</td></tr></tbody></table>

brain membrane fractionation: an ex vivo approach to assess subsynaptic protein localization

Assessing the synaptic protein composition and function constitutes an important challenge in neuroscience. However, it is not easy to evaluate neurotransmission that occurs within synapses because it is highly regulated by dynamic protein-protein interactions and phosphorylation events. Accordingly, when any method is used to study synaptic transmission, a major goal is to preserve these transient physiological modifications. Here, we present a brain membrane fractionation protocol that represents a robust procedure to isolate proteins belonging to different synaptic compartments. In other words, the protocol describes a biochemical methodology to carry out protein enrichment from presynaptic, postsynaptic, and extrasynaptic compartments. First, synaptosomes, or synaptic terminals, are obtained from neurons that contain all synaptic compartments by means of a discontinuous sucrose gradient. Of note, the quality of this initial synaptic membrane preparation is critical. Subsequently, the isolation of the different subsynaptic compartments is achieved with light solubilization using mild detergents at differential pH conditions. This allows for separation by gradient and isopycnic centrifugations. Finally, protein enrichment at the different subsynaptic compartments (i.e., pre-, post- and extrasynaptic membrane fractions) is validated by means of immunoblot analysis using well-characterized synaptic protein markers (i.e., SNAP-25, PSD-95, and synaptophysin, respectively), thus enabling a direct assessment of the synaptic distribution of any particular neuronal protein.

The described methodology has been largely used for the subsynaptic analysis of neuronal proteins in general and for the isolation and biochemical characterization of synaptic receptors5,6,7,8,9 in particular. Interestingly, the representative result displayed here show the usefulness of this experimental procedure for the anal...

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Brain Membrane Fractionation: An Ex Vivo Approach to Assess Subsynaptic Protein Localization

Here, we present a brain membrane fractionation protocol that represents a robust procedure to isolate proteins belonging to different synaptic compartments.

Brain Membrane Fractionation: An Ex Vivo Approach to Assess Subsynaptic Protein Localization

Assessing the synaptic protein composition and function constitutes an important challenge in neuroscience. However, it is not easy to evaluate ...

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Research

JoVE Journal

Biochemistry

10.3K Views.  Universitat de Barcelona, L'Hospitalet de Llobregat.  Hier presenteren wij een breukmembraan fractioneringsprotocol dat een robuuste procedure voor het isoleren van eiwitten uit verschillende synaptische compartimenten vertegenwoordigt. 

Video: Brain Membrane Fractionation: An Ex Vivo Approach to Assess Subsynaptic Protein Localization

Preparation of Synaptoneurosomes from Mouse Cortex using a Discontinuous Percoll-Sucrose Density Gradient

Synaptoneurosomes (SNs) are obtained after homogenization and fractionation of mouse brain cortex. They are resealed vesicles or isolated terminals that break away from axon terminals when the cortical tissue is homogenized. The SNs retain pre- and postsynaptic characteristics, which makes them useful in the study of synaptic transmission. They retain the molecular machinery used in neuronal signaling and are capable of uptake, storage, and release of neurotransmitters.
The production and isolation of active SNs can be problematic using medias like Ficoll, which can be cytotoxic and require extended centrifugation due to high density, and filtration and centrifugation methods, which can result in low activity due to mechanical damage of the SNs. However, the use of discontinuous Percoll-sucrose density gradients to isolate SNs provides a rapid method to produce good yields of translationally active SNs. The Percoll-sucrose gradient method is quick and gentle as it employs isotonic conditions, has fewer and shorter centrifugation spins and avoids centrifugation steps that pellet SNs and cause mechanical damage.

A method to prepare translationally active, intact synaptoneurosomes (SNs) from mouse brain cortex is described. The method uses a discontinuous Percoll-sucrose density gradient allowing for the quick preparation of active SNs.

Synaptoneurosomes (SNs) are obtained after homogenization and fractionation of mouse brain cortex. They are resealed vesicles or isolated terminals that ...

Neuroscience

Brain Slice Biotinylation: An Ex Vivo Approach to Measure Region-specific Plasma Membrane Protein Trafficking in Adult Neurons

Regulated endocytic trafficking is the central mechanism facilitating a variety of neuromodulatory events, by dynamically controlling receptor, ion channel, and transporter cell surface presentation on a minutes time scale.&#160;There is a broad diversity of mechanisms that control endocytic trafficking of individual proteins. Studies investigating the molecular underpinnings of trafficking have primarily relied upon surface biotinylation to quantitatively measure changes in membrane protein surface expression in response to exogenous stimuli and gene manipulation.&#160;However, this approach has been mainly limited to cultured cells, which may not faithfully reflect the physiologically relevant mechanisms at play in adult neurons.&#160;Moreover, cultured cell approaches may underestimate region-specific differences in trafficking mechanisms.&#160;Here, we describe an approach that extends cell surface biotinylation to the acute brain slice preparation.&#160;We demonstrate that this method provides a high-fidelity approach to measure rapid changes in membrane protein surface levels in adult neurons.&#160;This approach is likely to have broad utility in the field of neuronal endocytic trafficking.

Brain Slice Biotinylation: An Ex Vivo Approach to Measure Region-specific Plasma Membrane Protein Trafficking in Adult Neurons

Neuronal membrane trafficking dynamically controls plasma membrane protein availability and significantly impacts neurotransmission.&#160;To date, it has been challenging to measure neuronal endocytic trafficking in adult neurons.&#160;Here, we describe a highly effective, quantitative method to measure rapid changes in surface protein expression ex vivo in acute brain slices.

Regulated endocytic trafficking is the central mechanism facilitating a variety of neuromodulatory events, by dynamically controlling receptor, ion ...

Preparation of Synaptic Plasma Membrane and Postsynaptic Density Proteins Using a Discontinuous Sucrose Gradient

Neuronal subcellular fractionation techniques allow the quantification of proteins that are trafficked to and from the synapse. As originally described in the late 1960&#8217;s, proteins associated with the synaptic plasma membrane can be isolated by ultracentrifugation on a sucrose density gradient. Once synaptic membranes are isolated, the macromolecular complex known as the post-synaptic density can be subsequently isolated due to its detergent insolubility. The techniques used to isolate synaptic plasma membranes and post-synaptic density proteins remain essentially the same after 40 years, and are widely used in current neuroscience research. This article details the fractionation of proteins associated with the synaptic plasma membrane and post-synaptic density using a discontinuous sucrose gradient. Resulting protein preparations are suitable for western blotting or 2D DIGE analysis.

This article details the enrichment of proteins associated with the synaptic plasma membrane by ultracentrifugation on a discontinuous sucrose gradient. The subsequent preparation of post-synaptic density proteins is also described. Protein preparations are suitable for western blotting or 2D DIGE analysis.

Neuronal subcellular fractionation techniques allow the quantification of proteins that are trafficked to and from the synapse. As originally described in ...

In situ Subcellular Fractionation of Adherent and Non-adherent Mammalian Cells

Protein function is intimately coupled to protein localization. Although some proteins are restricted to a specific location or subcellular compartment, many proteins are present as a freely diffusing population in free exchange with a sub-population that is tightly associated with a particular subcellular domain or structure. In situ subcellular fractionation allows the visualization of protein compartmentalization and can also reveal protein sub-populations that localize to specific structures. For example, removal of soluble cytoplasmic proteins and loosely held nuclear proteins can reveal the stable association of some transcription factors with chromatin. Subsequent digestion of DNA can in some cases reveal association with the network of proteins and RNAs that is collectively termed the nuclear scaffold or nuclear matrix. 
Here we describe the steps required during the in situ fractionation of adherent and non-adherent mammalian cells on microscope coverslips. Protein visualization can be achieved using specific antibodies or fluorescent fusion proteins and fluorescence microscopy. Antibodies and/or fluorescent dyes that act as markers for specific compartments or structures allow protein localization to be mapped in detail. In situ fractionation can also be combined with western blotting to compare the amounts of protein present in each fraction. This simple biochemical approach can reveal associations that would otherwise remain undetected.

In situ Subcellular Fractionation of Adherent and Non-adherent Mammalian Cells

In situ subcellular fractionation of mammalian cells on microscope coverslips allows the visualisation of protein localisation.

Protein function is intimately coupled to protein localization. Although some proteins are restricted to a specific location or subcellular compartment, ...

Biology

Utilizing Combined Methodologies to Define the Role of Plasma Membrane Delivery During Axon Branching and Neuronal Morphogenesis

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular guidance cues like netrin-1 and results in dramatic increases to the surface area of the axonal plasma membrane. Netrin-1-dependent axon branching likely involves temporal and spatial control of plasma membrane expansion, the components of which are supplied through exocytic vesicle fusion. These fusion events are preceded by formation of SNARE complexes, comprising a v-SNARE, such as VAMP2 (vesicle-associated membrane protein 2), and plasma membrane t-SNAREs, syntaxin-1 and SNAP25 (synaptosomal-associated protein 25). Detailed herein isa multi-pronged approach used to examine the role of SNARE mediated exocytosis in axon branching. The strength of the combined approach is data acquisition at a range of spatial and temporal resolutions, spanning from the dynamics of single vesicle fusion events in individual neurons to SNARE complex formation and axon branching in populations of cultured neurons. This protocol takes advantage of established biochemical approaches to assay levels of endogenous SNARE complexes and Total Internal Reflection Fluorescence (TIRF) microscopy of cortical neurons expressing VAMP2 tagged with a pH-sensitive GFP (VAMP2-pHlourin) to identify netrin-1 dependent changes in exocytic activity in individual neurons. To elucidate the timing of netrin-1-dependent branching, time-lapse differential interference contrast (DIC) microscopy of single neurons over the order of hours is utilized. Fixed cell immunofluorescence paired with botulinum neurotoxins that cleave SNARE machinery and block exocytosis demonstrates that netrin-1 dependent axon branching requires SNARE-mediated exocytic activity.

Light microscopy techniques coupled with biochemical assays elucidate the involvement of SNARE-mediated exocytosis in netrin-dependent axon branching. This combination of techniques permits identification of molecular mechanisms controlling axon branching and cell shape change.

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular ...

High Resolution Quantitative Synaptic Proteome Profiling of Mouse Brain Regions After Auditory Discrimination Learning

The molecular synaptic mechanisms underlying auditory learning and memory remain largely unknown. Here, the workflow of a proteomic study on auditory discrimination learning in mice is described. In this learning paradigm, mice are trained in a shuttle box Go/NoGo-task to discriminate between rising and falling frequency-modulated tones in order to avoid a mild electric foot-shock. The protocol involves the enrichment of synaptosomes from four brain areas, namely the auditory cortex, frontal cortex, hippocampus, and striatum, at different stages of training. Synaptic protein expression patterns obtained from trained mice are compared to na&#239;ve controls using a proteomic approach. To achieve sufficient analytical depth, samples are fractionated in three different ways prior to mass spectrometry, namely 1D SDS-PAGE/in-gel digestion, in-solution digestion and phospho-peptide enrichment. 
High-resolution proteomic analysis on a mass spectrometer and label-free quantification are used to examine synaptic protein profiles in phospho-peptide-depleted and phospho-peptide-enriched fractions of synaptosomal protein samples. A commercial software package is utilized to reveal proteins and phospho-peptides with significantly regulated relative synaptic abundance levels (trained/na&#239;ve controls). Common and differential regulation modes for the synaptic proteome in the investigated brain regions of mice after training were observed. Subsequently, meta-analyses utilizing several databases are employed to identify underlying cellular functions and biological pathways.

The identification of molecules and pathways controlling synaptic plasticity and memory is still a major challenge in neuroscience. Here, a workflow is described addressing the relative quantification of synaptic proteins supposedly involved in the molecular reorganization of synapses during learning and memory consolidation in an auditory learning paradigm.

The molecular synaptic mechanisms underlying auditory learning and memory remain largely unknown. Here, the workflow of a proteomic study on auditory ...

Electroconvulsive Seizures in Rats and Fractionation of Their Hippocampi to Examine Seizure-induced Changes in Postsynaptic Density Proteins

Electroconvulsive seizure (ECS) is an experimental animal model of electroconvulsive therapy, the most effective treatment for severe depression. ECS induces generalized tonic-clonic seizures with low mortality and neuronal death and is a widely-used model to screen anti-epileptic drugs. Here, we describe an ECS induction method in which a brief 55-mA current is delivered for 0.5 s to male rats 200 - 250 g in weight via ear-clip electrodes. Such bilateral stimulation produced stage 4 - 5 clonic seizures that lasted about 10 s. After the cessation of acute or chronic ECS, most rats recovered to be behaviorally indistinguishable from sham &#34;no seizure&#34; rats. Because ECS globally elevates brain activity, it has also been used to examine activity-dependent alterations of synaptic proteins and their effects on synaptic strength using multiple methods. In particular, subcellular fractionation of the postsynaptic density (PSD) in combination with Western blotting allows for the quantitative determination of the abundance of synaptic proteins at this specialized synaptic structure. In contrast to a previous fractionation method that requires large amount of rodent brains, we describe here a small-scale fractionation method to isolate the PSD from the hippocampi of a single rat, without sucrose gradient centrifugation. Using this method, we show that the isolated PSD fraction contains postsynaptic membrane proteins, including PSD95, GluN2B, and GluA2. Presynaptic marker synaptophysin and soluble cytoplasmic protein &#945;-tubulin were excluded from the PSD fraction, demonstrating successful PSD isolation. Furthermore, chronic ECS decreased GluN2B expression in the PSD, indicating that our small-scale PSD fractionation method can be applied to detect the changes in hippocampal PSD proteins from a single rat after genetic, pharmacological, or mechanical treatments.

Electroconvulsive seizure (ECS) is an experimental animal model of electroconvulsive therapy for severe depression. ECS globally stimulates activity in the hippocampus, leading to synaptogenesis and synaptic plasticity. Here, we describe methods for ECS induction in rats and for subcellular fractionation of their hippocampi to examine seizure-induced changes in synaptic proteins.

Electroconvulsive seizure (ECS) is an experimental animal model of electroconvulsive therapy, the most effective treatment for severe depression. ECS ...

Quantifying the Heterogeneous Distribution of a Synaptic Protein in the Mouse Brain Using Immunofluorescence

The presence, absence, or levels of specific synaptic proteins can severely influence synaptic transmission. In addition to elucidating the function of a protein, it is vital to also determine its distribution. Here, we describe a protocol employing immunofluorescence, confocal microscopy, and computer-based analysis to determine the distribution of the synaptic protein Mover (also called TPRGL or SVAP30). We compare the distribution of Mover to that of the synaptic vesicle protein synaptophysin, thereby determining the distribution of Mover in a quantitative manner relative to the abundance of synaptic vesicles. Notably, this method could potentially be implemented to allow for comparison of the distribution of proteins using different antibodies or microscopes or across different studies. Our method circumvents the inherent variability of immunofluorescent stainings by yielding a ratio rather than absolute fluorescence levels. Additionally, the method we describe enables the researcher to analyze the distribution of a protein on different levels: from whole brain slices to brain regions to different subregions in one brain area, such as the different layers of the hippocampus or sensory cortices. Mover is a vertebrate-specific protein that is associated with synaptic vesicles. With this method, we show that Mover is heterogeneously distributed across brain areas, with high levels in the ventral pallidum, the septal nuclei, and the amygdala, and also within single brain areas, such as the different layers of the hippocampus.

Here, we describe a quantitative approach to determining the distribution of a synaptic protein relative to a marker protein using immunofluorescence staining, confocal microscopy, and computer-based analysis.

The presence, absence, or levels of specific synaptic proteins can severely influence synaptic transmission. In addition to elucidating the function of a ...

Subcellular Fractionation for the Isolation of Synaptic Components from the Murine Brain

Synaptic terminals are the primary sites of neuronal communication. Synaptic dysfunction is a hallmark of many neuropsychiatric and neurological disorders. The characterization of synaptic sub-compartments by biochemical isolation is, therefore, a powerful method to elucidate the molecular bases of synaptic processes, both in health and disease. This protocol describes the isolation of synaptic terminals and synaptic sub-compartments from mouse brains by subcellular fractionation. First, sealed synaptic terminal structures, known as synaptosomes, are isolated following brain tissue homogenization. Synaptosomes are neuronal pre- and post-synaptic compartments with pinched-off and sealed membranes. These structures retain a metabolically active state and are valuable for studying synaptic structure and function. The synaptosomes are then subjected to hypotonic lysis and ultracentrifugation to obtain synaptic sub-compartments enriched for synaptic vesicles, synaptic cytosol, and synaptic plasma membrane. Fraction purity is confirmed by electron microscopy and biochemical enrichment analysis for proteins specific to sub-synaptic compartments. The presented method is a straightforward and valuable tool for studying the structural and functional characteristics of the synapse and the molecular etiology of various brain disorders.

This protocol presents a robust, detailed method to obtain highly pure synaptosomes, synaptic vesicles, and other synaptic fractions from the mouse brain. This method enables the evaluation of synaptic processes, including the biochemical analysis of protein localization and function with compartmental resolution.

Synaptic terminals are the primary sites of neuronal communication. Synaptic dysfunction is a hallmark of many neuropsychiatric and neurological ...

Determination of Plasma Membrane Partitioning for Peripherally-associated Proteins

This method provides a fast approach for the determination of plasma membrane partitioning of any fluorescently-tagged peripherally-associated protein using the profiles of fluorescence intensity across the plasma membrane. Measured fluorescence profiles are fitted by a model for membrane and cytoplasm fluorescence distribution along a line applied perpendicularly to the cell periphery. This model is constructed from the fluorescence intensity values in reference cells expressing a fluorescently-tagged marker for cytoplasm and with FM 4-64-labeled plasma membrane. The method can be applied to various cell types and organisms; however, only plasma membranes of non-neighboring cells can be evaluated. This fast microscopy-based method is suitable for experiments, where subtle and dynamic changes of plasma membrane-associated markers are expected and need to be quantified, e.g., in the analysis of mutant versions of proteins, inhibitor treatments, and signal transduction observations. The method is implemented in a multi-platform R package that is coupled with an ImageJ macro that serves as a user-friendly interface.

Here, we present a protocol to perform a quantitative analysis of the level of plasma-membrane association for fluorescently-tagged peripherally-associated protein. The method is based on the computational decomposition of membrane and cytoplasmic component of signal observed in cells labeled with plasma membrane fluorescent marker.

This method provides a fast approach for the determination of plasma membrane partitioning of any fluorescently-tagged peripherally-associated protein ...

Brain Membrane Fractionation: An Ex Vivo Approach to Assess Subsynaptic Protein Localization

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