This work was supported by startup funds from the College of Agricultural and Life Sciences and the College of Science at Virginia Tech. T.M. is supported by the George Washington Carver Program at Virginia Tech.

All procedures including animal subjects have been approved by the Virginia Polytechnic Institute and State University Institutional Animal Care and Use Committee (IACUC).
1. Collection and Dissection of Rodent Brain Tissue
NOTE: This protocol can be applied to a variety of brain regions and used with various tissue collection procedures. Below is the procedure used in our lab for subcellular of rat brain tissue, using 8-9 week old male Sprague Dawley rats. In order to process all cellular compartments in the same animal, section 1.3. must be followed regardless of the tissue collection procedure used.
<ol>
	<li>Extract the rat brain and place into a cryogenic cup pre-chilled on dry ice. Flash-freeze the brain on dry ice, or use liquid nitrogen if available, and transfer to a -80 &#176;C freezer. Brains can be dissected the same day or at a later time.</li>
	<li>Remove the frozen brain from the cryogenic cup and place into a rat brain matrix chilled with dry ice. Each slot on the matrix corresponds to 0.5 mm, which can be used to determine the approximate location of the region of interest (ROI).
	<ol>
		<li>Using a Rat Brain Atlas, insert a razor blade into the matrix immediately in front (anterior) of the predicted start of ROI. Next, insert a razor blade immediately at the predicted end (posterior) of ROI.</li>
		<li>Remove the slice between the razor blades using a scalpel and place onto a microscope slide chilled on dry ice. Typically, sections are 2-3 mm thick but this varies based on the ROI.</li>
	</ol>
	</li>
	<li>Using a scalpel, dissect out the ROI one hemisphere at a time. Place each hemisphere into separate 1.5 mL microcentrifuge tubes pre-chilled on dry ice. Frozen tissue can immediately be used for subcellular fractionation or processed at a later time if stored at -80 &#176;C.</li>
</ol>
2. Nuclear and Cytoplasmic Extraction
NOTE: This protocol uses premade stock solutions of common lab chemicals, including 0.1 M HEPES, 1 M MgCl2, 1 M Dithiothreitol (DTT), 0.5 M ethylenediaminetetraacetic acid (EDTA), 5 M NaCl, 10% NP-40 (IGEPAL) and 50% glycerol. If endpoint is used in proteasome activity assays, glycerol and ATP can be added to all buffers to help prevent disassembly of proteasome complexes during lysis.
<ol>
	<li>Prepare the Lysis Buffer. Add 8.63 mL of ultrapure (deionized and distilled) water to a sterile 15 mL conical tube.
	<ol>
		<li>To the conical tube, add 1,000 &#181;L of 0.1 M HEPES, 15 &#181;L of 1 M MgCl, 100 &#181;L of 1 M DTT and 50 &#181;L of 10% NP-40.</li>
		<li>Add 100 &#181;L of protease inhibitor cocktail and 100 &#181;L of phosphatase inhibitor cocktail. Briefly vortex the solution to mix and place on wet ice to chill. Note that the solution may have a yellow tint from the phosphatase inhibitor. 
		NOTE: The use of protease inhibitors is essential to preserve protein and ensure the specificity of the in vitro&#160;proteasome activity assay. However, these inhibitors may also result in a slight reduction in proteasome activity, meaning that the activity measured on the in vitro assay may be an underestimate of actual activity levels.</li>
	</ol>
	</li>
	<li>Prepare the Extraction Buffer. Add 5.925 mL of ultrapure (deionized and distilled) water to a sterile 15 mL conical tube.
	<ol>
		<li>To the conical tube, add 2,000 &#181;L of 0.1 M HEPES, 1250 &#181;L of 50% glycerol, 15 &#181;L of 1 M MgCl2, 5 &#181;L of 1 M DTT, 5 &#181;L of 0.5 M EDTA and 600 &#181;L of 5 M NaCl.</li>
		<li>Add 100 &#181;L of protease inhibitor cocktail and 100 &#181;L of phosphatase inhibitor cocktail. Briefly vortex the solution to mix and place on wet ice to chill. Note that the solution may have a yellow tint from the phosphatase inhibitor.</li>
	</ol>
	</li>
	<li>Remove the 1.5 mL centrifuge microtube containing one hemisphere of the ROI from the -80 &#176;C freezer. Ensure that the hemisphere used is counterbalanced across extraction conditions for each experimental group. For example, in a two-group experiment, use the left hemisphere for the first two animals in each group and the right hemisphere for the next two samples, etc.</li>
	<li>Using a scalpel, transfer the frozen brain tissue to a 2 mL glass Teflon homogenizer. Add 500 &#181;L of Lysis buffer to the Teflon tube.
	<ol>
		<li>Using pestle B, homogenize the same tissue using 15 strokes until no visible amount of solid material are present. Use a turning motion (clockwise or counter-clockwise) during each stroke.</li>
	</ol>
	</li>
	<li>Using a 1,000 &#181;L pipette, transfer the homogenized sample to a new 1.5 mL microcentrifuge tube. Place the tube on wet ice and incubate for 30 min.</li>
	<li>Place tube in microcentrifuge and spin for 10 min at 845 x g and 4 &#176;C. After completion, carefully remove the supernatant by pipetting and place into a new 1.5 mL microcentrifuge tube; this is the Cytoplasmic Fraction, which can be stored on ice or at 4 &#176;C until Section 2.9.</li>
	<li>Add 50 &#181;L of Extraction buffer to the resulting pellet and resuspend by pipetting. Do not vortex the pellet. Place the tube containing the resuspended pellet on ice and incubate for 30 min.</li>
	<li>Place the tube in microcentrifuge and spin for 20 min at 21,456 x g, 4 &#176;C. After completion, carefully remove the supernatant by pipetting and place into a new 1.5 mL microcentrifuge tube; this is the Nuclear Fraction. The pellet can now be discarded.</li>
	<li>Measure protein concentration of the Cytoplasmic and Nuclear extractions using the Dc protein assay (according to the manufacturer&#39;s instructions), or any equivalent assay for samples collected using non-ionic detergents. Proceed immediately to proteasome activity assay (Section 4) or Western blotting (Section 5). Alternatively, samples can be stored at -80 &#176;C until needed.</li>
</ol>
3. Synaptic Fraction Collection
NOTE: This protocol uses premade stock solutions of common lab chemicals, including 1 M Tris (pH 7.5), 0.5 M EDTA, 5 M NaCl and 10% SDS. If endpoint is used in proteasome activity assays, glycerol and ATP can be added to all buffers to help prevent disassembly of proteasome complexes during lysis
<ol>
	<li>Prepare the TEVP buffer with 320 mM sucrose. Add 60 mL of ultrapure (deionized and distilled) water to a clean 100 mL beaker.
	<ol>
		<li>To the beaker, add 1,300 &#181;L of 1 M Tris (pH 7.5), 260 &#181;L of 0.5 M EDTA, 100 &#181;L of protease inhibitor cocktail, 100 &#181;L of phosphatase inhibitor cocktail and 10.944 g of sucrose. Mix with a stir bar until sucrose is completely dissolved.</li>
		<li>Bring pH to ~7.4 by adding hydrochloric acid (HCl) dropwise to the solution using a pipette.</li>
		<li>Transfer the solution to a 100 mL volumetric flask. Bring volume to 100 mL by adding ultrapure water. Chill the final solution on ice until needed.</li>
	</ol>
	</li>
	<li>Prepare the Homogenization Buffer. Add 60 mL of ultrapure (deionized and distilled) water to a clean 100 mL beaker.
	<ol>
		<li>To the beaker, add 5,000 &#181;L of 1 M Tris (pH 7.5), 3,000 &#181;L of 5 M NaCl, 100 &#181;L of protease inhibitor cocktail, 100 &#181;L of phosphatase inhibitor cocktail and 2,000 &#181;L of 10% SDS. Mix with a stir bar. 
		NOTE: Ionic detergents can interfere with proteasome activity assays by resulting in denaturing of the proteasome complex. The volume of SDS could be lowered to help preserve proteasome function if desired.</li>
		<li>Bring pH to ~7.4 by adding HCl dropwise to the solution using a pipette.</li>
		<li>Transfer the solution to a 100 mL volumetric flask. Bring the volume to 100 mL by adding ultrapure water. Store the final solution at room temperature; do not chill as SDS will precipitate out of solution.</li>
	</ol>
	</li>
	<li>Remove the 1.5 mL centrifuge containing one hemisphere of the ROI from the -80 &#176;C freezer. Ensure that the hemisphere used is counterbalanced across extraction conditions for each experimental group. For example, in a two-group experiment, use the left hemisphere for the first two animals in each group and the right hemisphere for the next two samples, etc.</li>
	<li>Using a scalpel, transfer the frozen brain tissue to a 2 mL glass Teflon homogenizer. Add 500 &#181;L of TEVP buffer to the Teflon tube.
	<ol>
		<li>Using pestle B, homogenize the same tissue using 15 strokes until no visible transfers of solid material are present. Use a turning motion (clockwise or counter-clockwise) during each stroke.</li>
	</ol>
	</li>
	<li>Using a 1,000 &#181;L pipette, transfer the homogenized sample to a new 1.5 mL microcentrifuge tube. Centrifuge the sample at 1,000 x g for 10 min, 4 &#176;C.</li>
	<li>Collect the supernatant and transfer to a new 1.5 mL microcentrifuge tube using a 1,000 &#181;L pipette. Centrifuge the sample at 10,000 x g for 10 min, 4 &#176;C. The original pellet (P1) contains nuclei and the large debris and can be discarded.</li>
	<li>Transfer the supernatant to a new 1.5 mL microcentrifuge tube. This is a Cytosolic Fraction. Add 50 &#181;L of Homogenization buffer to the pellet (P2) and resuspend by pipetting until no solid material is visible.</li>
	<li>Centrifuge the sample at 20,000 x g for 10 min, 4 &#176;C. Transfer the supernatant to a new 1.5 mL microcentrifuge tube; this is the Crude Synaptosomal Membrane (Synaptic) Fraction. The pellet can be discarded.</li>
	<li>Measure the protein concentration of the Synaptic fraction using the Dc protein assay (according to the manufacturer&#39;s instructions), or any equivalent assay for samples collected using ionic detergents. Proceed immediately to proteasome activity assay (Section 4) or Western blotting (Section 5). Alternatively, samples can be stored at -80 &#176;C until needed.</li>
</ol>
4. Proteasome Activity Assay
NOTE: Proteasome activity can be measured in homogenized brain tissue using a slightly modified version of the 20S Proteasome Activity Kit. This assay does not directly measure the activity of complete 26S proteasome complexes. Rather, it measures the activity of the 20S core, meaning it can only serve as a proxy to understand the activity of the core itself as opposed to the entire 26S proteasome complex. The success of this assay declines with repeated freeze-thaw cycles and/or increasing levels of detergents, particularly ionic, and requires the use of a plate reader with a 360/460 (excitation/emission) filter set and heating capabilities up to 37 &#176;C.
<ol>
	<li>Plate reader settings: Pre-warm to 37 &#176;C and hold through the run.
	<ol>
		<li>Set excitation to 360 and emission to 460. If the 96 well plate used is clear, set the optics position to Bottom. If a dark/black 96 well plate is used, set optics position to Top.</li>
		<li>Set older plate reader models to Auto-Gain under the fluorescent read options; newer models are preset for this. Program a kinetic run with a time of 2 h, scanning (reading) every 30 min.</li>
	</ol>
	</li>
	<li>Reconstitute the 10x assay buffer provided in the kit with 13.5 mL of ultrapure water. Add 14 &#181;L of 100 mM ATP to the now 1x buffer; this significantly enhances proteasome activity in the samples and improves assay reliability. The final 20S Assay buffer can be stored on ice or at 4 &#176;C until needed and is stable for several months.</li>
	<li>Reconstitute the AMC standard provided in kit with 100 &#181;L of DMSO. Perform this step in the dark or under low light conditions, as the standard is light sensitive.</li>
	<li>Create a stand curve of AMC using the reconstituted standard, in the dark or under low light conditions.
	<ol>
		<li>In separate 0.5 mL microcentrifuge tubes, add 16, 8, 6.4, 3.2, 1.6, 0.8, 0.4 and 0 &#181;L of the AMC standard, which corresponds to 20, 10, 8, 4, 2, 1, 0.5 and 0 &#181;M AMC concentration.</li>
		<li>To these tubes, in the same order, add 84, 92, 93.6, 96.8, 98.4, 99.2, 99.6 and 100 &#181;L of 20S Assay buffer. This creates a series of high to low AMC concentrations, which will be used for plate reader calibration and analysis of proteasome activity in the homogenized samples.</li>
		<li>Store all diluted standards on ice in the dark until needed.</li>
	</ol>
	</li>
	<li>Reconstitute the proteasome substrate (Suc-LLVY-AMC) provided in kit with 65 &#181;L of DMSO. Perform this step in the dark or under low light conditions, as the substrate is light sensitive. Create a 1:20 dilution of the proteasome substrate in a new 1.5 mL microcentrifuge tube using 20S Assay buffer. Store the diluted substrate on ice in the dark until needed. 
	NOTE: For example, if the plate will have 10 samples and 1 blank, you will need enough diluted substrate for 22 wells (with duplicates) at 10 &#181;L per well. This equates to 220 &#181;L need + 30 &#181;L for pipetting errors, requiring 12.5 &#181;L of substrate and 237.5 &#181;L of 20S Assay buffer.</li>
	<li>Thaw desired samples (if frozen) and add a normalized amount to a 96 well plate. Run each sample in duplicates. The amount of sample needed varies based on tissue preparation. Generally, 10-20 &#181;g is sufficient for any subcellular fraction.</li>
	<li>Bring the sample well volume to 80 &#181;L with ultrapure water. The amount added depends on the volume of sample added. For example, if sample 1 was 4.5 &#181;L of protein and sample 2 was 8.7 &#181;L, then the amount of water needed will be 75.5 &#181;L and 71.3 &#181;L, respectively. In two separate wells, add 80 &#181;L of water alone; these will be the Assay Blanks. 
	NOTE: In order to limit changes in protein volume due to pipetting errors, protein concentrations can be normalized to the least concentrated sample. This will allow use of the same sample volume in all conditions.</li>
	<li>Add 10 &#181;L of 20S Assay buffer to each well, including Assay Blanks. A repeater/automated pipette is recommended here to ensure consistent assay volume across wells.</li>
	<li>Optional: At this stage, introduce in vitro&#160;manipulations if desired; this will require that each sample has an additional 2 wells per treatment, including the vehicle. If so, add 5-10 &#181;L of drug/compound of interest to 2 of the sample wells and an equivalent volume of control/vehicle to another 2 wells. Place the plate onto the pre-warmed plate reader or into a 37 &#176;C incubator for 30 min.</li>
	<li>Turn off the lights or enter a dark room. Add all 100 &#181;L of diluted AMC standards to a new well; each standard will have a single well.</li>
	<li>In the dark, add 10 &#181;L of diluted proteasome substrate to wells containing sample and Assay Blanks, but not to the AMC standard. A repeater/automated pipette is recommended here to ensure consistent assay volume across wells.</li>
	<li>Place the plate into the plate reader and start the kinetic run. 
	NOTE: The plate does not need to be under constant agitation during the kinetic run, however, the user can choose to if desired</li>
	<li>At end of the kinetic run, export raw 360/460 fluorescent values to Microsoft Excel.
	<ol>
		<li>Average together duplicate wells for each standard, each sample and the Assay Blank for all 5 scans. Raw fluorescent values should increase across scans for the samples but remain stable (or decrease slightly) for standards and Assay Blanks.</li>
		<li>Take the highest AMC standard well average and divide by the known concentration (20 &#181;M). Divide this value by the sample concentration used in the assay to get standardized AMC value. For each sample and Assay Blank average, divide by the standardized AMC value.</li>
		<li>Take this final value and divide by the sample concentration used in the assay to get the normalized value for each sample and the Assay Blank. Do this for all 5 scans.</li>
		<li>Subtract the normalized Assay Blank value from each normalized sample value for all 5 scans.</li>
	</ol>
	</li>
</ol>
5. Quantification of Linkage-specific Protein Ubiquitination
<ol>
	<li>Quantification of diverse polyubiquitin tags in different subcellular fractions collected from rodent brain tissue should be done using a variety of standard Western blotting protocols in combination with unique, linkage-specific polyubiquitin antibodies.
	<ol>
		<li>For denaturing, mix the normalized samples with an equal volume of Laemmli sample buffer supplemented with &#946;-mercaptoethanol to a percentage of 5% by volume, per the manufacturer&#39;s instruction.</li>
		<li>For quantification of all monoubiquitination and polyubiquitination modifications regardless of linkage, use a pan-ubiquitin antibody.</li>
		<li>To recognize all polyubiquitinated proteins, use an ubiquitin antibody that does not cross-react with monoubiquitination.</li>
		<li>For linkage-specific polyubiquitination, use antibodies that can detect Lysine-27, Lysine-48, Lysine-63 and linear (M1) polyubiquitination.</li>
	</ol>
	</li>
	<li>To prevent cross-contamination between developments, which could result in a false positive or interfere with imaging of a different ubiquitin modification, strip membranes between developments using 0.1 M NaOH for 10 min.
	<ol>
		<li>Wash the membranes in TBS with 0.1% tween twice for 10 min and reblock (using whatever blocking agent is preferred in the Western blot protocol used).</li>
		<li>Incubate the membrane with the secondary antibody and redevelop in order to confirm that the membrane was stripped of primary and secondary antibodies properly.</li>
		<li>Recommended: For successful development of different ubiquitin antibodies without contamination, use fluorescent or near-infrared imaging systems. This often times can prevent carryover between antibodies.</li>
	</ol>
	</li>
	<li>Some ubiquitin Western blot images will provide columns of distinct bands (such as M1) while others produce a smear-like pattern with few or no clear lines (common with K48). For quantification of imaged ubiquitin Western blots, draw a box around the column that extends the entire molecular standards ladder.
	<ol>
		<li>Adjust the box up (or down) if ubiquitin staining does extend through the entire ladder; this is common for Lysine-48 modifications and varies widely across subcellular compartments.</li>
		<li>Subtract out the background, which is calculated as the mean optical density of the background immediately surrounding the column on all sides.</li>
	</ol>
	</li>
</ol>

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A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+subcellular+changes+in+proteasome+activity+and+linkage-specific+protein+polyubiquitination+in+the+amygdala+during+the+consolidation+and+reconsolidation+of+a+fear+memory.">Distinct subcellular changes in proteasome activity and linkage-specific protein polyubiquitination in the amygdala during the consolidation and reconsolidation of a fear memory.</a> Neurobiology of Learning and Memory. 157, 1-11 (2019).</li><li>Cullen, P. K., Ferrara, N. C., Pullins, S. E., Helmstetter, F. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Context+memory+formation+requires+activity-dependent+protein+degradation+in+the+hippocampus.">Context memory formation requires activity-dependent protein degradation in the hippocampus.</a> Learning and Memory. 24 (11), 589-596 (2017).</li><li>Jarome, T. J., Ferrara, N. C., Kwapis, J. L., Helmstetter, F. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CaMKII+regulates+proteasome+phosphorylation+and+activity+and+promotes+memory+destabilization+following+retrieval.">CaMKII regulates proteasome phosphorylation and activity and promotes memory destabilization following retrieval.</a> Neurobiology of Learning and Memory. 128, 103-109 (2016).</li><li>Lee, S. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synaptic+protein+degradation+underlies+destabilization+of+retrieved+fear+memory.">Synaptic protein degradation underlies destabilization of retrieved fear memory.</a> Science. 319 (5867), 1253-1256 (2008).</li><li>Dunah, A. W., Standaert, D. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dopamine+D1+receptor-dependent+trafficking+of+striatal+NMDA+glutamate+receptors+to+the+postsynaptic+membrane.">Dopamine D1 receptor-dependent trafficking of striatal NMDA glutamate receptors to the postsynaptic membrane.</a> Journal of Neuroscience. 21 (15), 5546-5558 (2001).</li><li>Kelly, P. T., Cotman, C. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synaptic+proteins.+Characterization+of+tubulin+and+actin+and+identification+of+a+distinct+postsynaptic+density+polypeptide.">Synaptic proteins. Characterization of tubulin and actin and identification of a distinct postsynaptic density polypeptide.</a> Journal of Cell Biology. 79 (1), 173-183 (1978).</li><li>Mengual, E., Arizti, P., Rodrigo, J., Gimenez-Amaya, J. M., Castano, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immunohistochemical+distribution+and+electron+microscopic+subcellular+localization+of+the+proteasome+in+the+rat+CNS.">Immunohistochemical distribution and electron microscopic subcellular localization of the proteasome in the rat CNS.</a> Journal of Neuroscience. 16 (20), 6331-6341 (1996).</li></ol>

The authors have nothing to disclose.

Here, we demonstrate an efficient method for quantifying changes in ubiquitin-proteasome activity across different subcellular compartments in the same animal. Currently, most attempts at measuring subcellular changes in activity of the ubiquitin-proteasome system have been limited to a single compartment per sample, resulting in the need to repeat experiments. This leads to significant costs and loss of animal life. Our protocol alleviates this problem by splitting hemispheres, allowing different cellular fractions to b...

The ubiquitin-proteasome system (UPS) is a complex network of interconnected protein structures and ligases that controls the degradation of most short-lived proteins in cells1. In this system, proteins are marked for degradation or other cellular processes/fates by the small modifier ubiquitin. A target protein can acquire 1-7 ubiquitin modifications, which can link together at one of seven lysine (K) sites (K6, K11, K27, K29, K33, K48 and K63) or the N-terminal methionine (M1; as known as linear) in the previous ubiquitin2. Some of these polyubiquitin tags are degradation-specific (K48)3, while others are largely independent of the protein degradation process (M1)4,5,6. Thus, the protein ubiquitination process is incredibly complex and the ability to quantify changes in a specific polyubiquitin tag is critical for ultimately understanding the role of that given modification in cellular functioning. Further complicating the study of this system, the proteasome, which is the catalytic structure of the UPS7, both degrades proteins but can also be involved in other non-proteolytic processes8,9. Not surprisingly then, since its initial discovery, normal and aberrant ubiquitin-proteasome activity has been implicated in long-term memory formation and a variety of disease states, including many neurological, neurodegenerative and psychiatric disorders10,11. As a result, methods which can effectively and efficiently quantify UPS activity in the brain are critical for ultimately understanding how this system is dysregulated in disease states and the eventual development of treatment options targeting ubiquitin and/or proteasome functioning.
There are a number of issues in quantifying ubiquitin-proteasome activity in brain tissue from rats and mice, which are the most common model systems used to study UPS function, including 1) the diversity of ubiquitin modifications, and 2) distribution and differential regulation of UPS functioning across subcellular compartments12,13,14. For example, many of the early demonstrations of ubiquitin-proteasome function in the brain during memory formation used whole cell lysates and indicated time-dependent increases in both protein ubiquitination and proteasome activity15,16,17,18,19,20. However, we recently found that ubiquitin-proteasome activity varied widely across subcellular compartments in response to learning, with simultaneous increases in some regions and decreases in others, a pattern that differs significantly from what was previously reported in whole cell lysates21. This is consistent with the limitation of a whole cell approach, as it cannot dissociate the contribution of changes in UPS activity across different subcellular compartments. Though more recent studies have employed synaptic fraction protocols to study the UPS specifically at synapses in response to learning22,23,24, the methods used occlude the ability to measure nuclear and cytoplasmic ubiquitin-proteasome changes in the same animal. This results in an unnecessary need to repeat experiments multiple times, collecting a different subcellular fraction in each. Not only does this result in a greater loss of animal lives, but it eliminates the ability to directly compare UPS activity across different subcellular compartments in response to a given event or during a specific disease state. Considering that protein targets of ubiquitin and the proteasome vary widely throughout the cell, understanding how ubiquitin-proteasome signaling differs in distinct subcellular compartments is critical for identifying the functional role of the UPS in the brain during memory formation and neurological, neurodegenerative and psychiatric disorders.
To address this need, we recently developed a procedure in which nuclear, cytoplasmic and synaptic fractions could be collected for a given brain region from the same animal21. Additionally, to account for the limited amount of protein that can be obtained from collecting multiple subcellular fractions from the same sample, we optimized previously established protocols to assay in vitro&#160;proteasome activity and linkage-specific protein ubiquitination in lysed cells collected from rodent brain tissue. Using this protocol, we were able to collect and directly compare learning-dependent changes in proteasome activity, K48, K63, M1 and overall protein polyubiquitination levels in the nucleus and cytoplasm and at synapses in the lateral amygdala of rats. Here, we describe in detail our procedure (Figure 1), which could significantly improve our understanding of how the UPS is involved in long-term memory formation and various disease states. However, it should be noted that the in vitro proteasome activity discussed in our protocol, while widely used, does not directly measure the activity of complete 26S proteasome complexes. Rather, this assay measures the activity of the 20S core, meaning it can only serve as a proxy to understand the activity of the core itself as opposed to the entire 26S proteasome complex.

<table border="0" cellpadding="0" cellspacing="0" style="width:878px;" width="878">
	<tbody>
		<tr height="40">
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			<td style="width:255px;">Various other vendors</td>
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			<td height="40" style="height:40px;">20S Proteasome Activity Kit</td>
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			<td>Other vendors carry different versions</td>
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			<td>Various other vendors</td>
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			<td>BioTek</td>
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			<td>Other vendors carry different versions</td>
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			<td>Fisher</td>
			<td>ICN19024280</td>
			<td>Various other vendors</td>
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			<td>Various other vendors</td>
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			<td>Varous other vendors</td>
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			<td style="width:255px;">Various other vendors</td>
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		<tr height="40">
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			<td>Abcam</td>
			<td>ab1791</td>
			<td>Various other vendors</td>
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			<td style="width:143px;">Millipore Sigma</td>
			<td style="width:259px;">H3375</td>
			<td style="width:255px;">Various other vendors</td>
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		<tr height="40">
			<td height="40" style="height:40px;">Hydrochloric acid</td>
			<td>Fisher</td>
			<td>SA48</td>
			<td>Various other vendors</td>
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			<td height="40" style="height:40px;width:222px;">IGEPAL (NP-40)</td>
			<td style="width:143px;">Millipore Sigma</td>
			<td style="width:259px;">I3021</td>
			<td style="width:255px;">Various other vendors</td>
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		<tr height="40">
			<td height="40" style="height:40px;">K48 Ubiquitin Antibody</td>
			<td>Abcam</td>
			<td>ab140601</td>
			<td>Various other vendors</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">K63 Ubiquitin Antibody</td>
			<td>Abcam</td>
			<td>ab179434</td>
			<td>Various other vendors</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:222px;">KCl</td>
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			<td style="width:259px;">S3014</td>
			<td style="width:255px;">Various other vendors</td>
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			<td>Various other vendors</td>
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		<tr height="40">
			<td height="40" style="height:40px;width:222px;">Protease Inhibitor</td>
			<td style="width:143px;">Millipore Sigma</td>
			<td style="width:259px;">P8340</td>
			<td style="width:255px;">Various other vendors</td>
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		<tr height="40">
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			<td>Cell signaling</td>
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			<td>Various other vendors</td>
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		<tr height="50">
			<td height="50" style="height:50px;width:222px;">SDS</td>
			<td style="width:143px;">Millipore Sigma</td>
			<td style="width:259px;">L3771</td>
			<td style="width:255px;">Various other vendors</td>
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			<td>SS255</td>
			<td>Various other vendors</td>
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			<td height="48" style="height:48px;width:222px;">Sucrose</td>
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			<td style="width:259px;">S0389</td>
			<td style="width:255px;">Various other vendors</td>
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			<td height="49" style="height:50px;width:222px;">Tris</td>
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			<td style="width:259px;">T1503</td>
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			<td>Various other vendors</td>
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		<tr height="48">
			<td height="48" style="height:48px;">Ubiquitin Antibody</td>
			<td>Enzo Life Sciences</td>
			<td>BML-PW8810</td>
			<td>Various other vendors</td>
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	</tbody>
</table>

quantifying subcellular ubiquitin-proteasome activity in the rodent brain

The ubiquitin-proteasome system is a key regulator of protein degradation and a variety of other cellular processes in eukaryotes. In the brain, increases in ubiquitin-proteasome activity are critical for synaptic plasticity and memory formation and aberrant changes in this system are associated with a variety of neurological, neurodegenerative and psychiatric disorders. One of the issues in studying ubiquitin-proteasome functioning in the brain is that it is present in all cellular compartments, in which the protein targets, functional role and mechanisms of regulation can vary widely. As a result, the ability to directly compare brain ubiquitin protein targeting and proteasome catalytic activity in different subcellular compartments within the same animal is critical for fully understanding how the UPS contributes to synaptic plasticity, memory and disease. The method described here allows collection of nuclear, cytoplasmic and crude synaptic fractions from the same rodent (rat) brain, followed by simultaneous quantification of proteasome catalytic activity (indirectly, providing activity of the proteasome core only) and linkage-specific ubiquitin protein tagging. Thus, the method can be used to directly compare subcellular changes in ubiquitin-proteasome activity in different brain regions in the same animal during synaptic plasticity, memory formation and different disease states. This method can also be used to assess the subcellular distribution and function of other proteins within the same animal.

Using the procedure described here, nuclear, cytoplasmic and synaptic fractions were collected from the lateral amygdala of the rat brain (Figure 1). Purity of the individual fractions were confirmed via Western blotting, probing with antibodies against proteins that should be enriched or depleted in the lysate. In the first hemisphere where a crude synaptic fraction was collected, postsynaptic density protein 95 (PSD95) was present in the synaptic, but not n...

Watch this Scientific Journal Video about Quantifying Subcellular Ubiquitin-proteasome Activity in the Rodent Brain at JoVE.com

Quantifying Subcellular Ubiquitin-proteasome Activity in the Rodent Brain

This protocol is designed to efficiently quantify ubiquitin-proteasome system (UPS) activity in different cellular compartments of the rodent brain. Users are able to examine UPS functioning in nuclear, cytoplasmic and synaptic fractions in the same animal, reducing the amount of time and number of animals needed to perform these complex analyses.

The ubiquitin-proteasome system is a key regulator of protein degradation and a variety of other cellular processes in eukaryotes. In the brain, increases ...

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6.6K Views.  Virginia Polytechnic Institute and State University. This protocol is designed to efficiently quantify ubiquitin-proteasome system (UPS) activity in different cellular compartments of the rodent brain. Users are able to examine UPS functioning in nuclear, cytoplasmic and synaptic fractions in the same animal, reducing the amount of time and number of animals needed to perform these complex analyses. 

Video: Quantifying Subcellular Ubiquitin-proteasome Activity in the Rodent Brain

Simultaneous fMRI and Electrophysiology in the Rodent Brain

To examine the neural basis of the blood oxygenation level dependent (BOLD) magnetic resonance imaging (MRI) signal, we have developed a rodent model in which functional MRI data and in vivo intracortical recording can be performed simultaneously. The combination of MRI and electrical recording is technically challenging because the electrodes used for recording distort the MRI images and the MRI acquisition induces noise in the electrical recording. To minimize the mutual interference of the two modalities, glass microelectrodes were used rather than metal and a noise removal algorithm was implemented for the electrophysiology data. In our studies, two microelectrodes were separately implanted in bilateral primary somatosensory cortices (SI) of the rat and fixed in place. One coronal slice covering the electrode tips was selected for functional MRI. Electrode shafts and fixation positions were not included in the image slice to avoid imaging artifacts. The removed scalp was replaced with toothpaste to reduce susceptibility mismatch and prevent Gibbs ringing artifacts in the images. The artifact structure induced in the electrical recordings by the rapidly-switching magnetic fields during image acquisition was characterized by averaging all cycles of scans for each run. The noise structure during imaging was then subtracted from original recordings. The denoised time courses were then used for further analysis in combination with the fMRI data. As an example, the simultaneous acquisition was used to determine the relationship between spontaneous fMRI BOLD signals and band-limited intracortical electrical activity. Simultaneous fMRI and electrophysiological recording in the rodent will provide a platform for many exciting applications in neuroscience in addition to elucidating the relationship between the fMRI BOLD signal and neuronal activity.

We have developed a method for simultaneous functional magnetic resonance imaging and electrophysiological recording in the rodent brain, providing a platform for the investigation of the relationship between neural activity and the blood oxygenation level dependent (BOLD) MRI signal.

To examine the neural basis of the blood oxygenation level dependent (BOLD) magnetic resonance imaging (MRI) signal, we have developed a rodent model in ...

Quantifying the Activity of cis-Regulatory Elements in the Mouse Retina by Explant Electroporation

Transcription factors within cellular gene networks control the spatiotemporal pattern and levels of expression of their target genes by binding to cis-regulatory elements (CREs), short (&tilde;300-600 bp) stretches of genomic DNA which can lie upstream, downstream, or within the introns of the genes they control. CREs (i.e., enhancers/promoters) typically consist of multiple clustered binding sites for both transcriptional activators and repressors1-3. They serve as logical integrators of transcriptional input giving a unitary output in the form of spatiotemporally precise and quantitatively exact promoter activity. Most studies of mammalian cis-regulation to date have relied on mouse transgenesis as a means of assaying the enhancer function of CREs4-5. This technique is time-consuming, costly and, on account of insertion site effects, largely non-quantitative. On the other hand, quantitative assays for mammalian CRE function have been developed in tissue culture systems (e.g., dual luciferase assays), but the in vivo relevance of these results is often uncertain. 
Electroporation offers an excellent alternative to traditional mouse transgenesis in that it permits both spatiotemporal and quantitative assessment of cis-regulatory activity in living mammalian tissue. This technique has been particularly useful in the analysis of cis-regulation in the central nervous system, especially in the cerebral cortex and the retina6-8. While mouse retinal electroporation, both in vivo and ex vivo, has been developed and extensively described by Matsuda and Cepko6-7,9, we have recently developed a simple approach to quantify the activity of photoreceptor-specific CREs in electroporated mouse retinas10. Given that the amount of DNA that is introduced into the retina by electroporation can vary from experiment to experiment, it is necessary to include a co-electroporated 'loading control' in all experiments. In this respect, the technique is very similar to the dual luciferase assay used to quantify promoter activity in cultured cells. 
When assaying photoreceptor cis-regulatory activity, electroporation is usually performed in newborn mice (postnatal day 0, P0) which is the time of peak rod production11-12. Once retinal cell types become post-mitotic, electroporation is much less efficient. Given the high rate of rod birth in newborn mice and the fact that rods constitute more than 70% of the cells in the adult mouse retina, the majority of cells that are electroporated at P0 are rods. For this reason, rod photoreceptors are the easiest retinal cell type to study via electroporation. The technique we describe here is primarily useful for quantifying the activity of photoreceptor CREs.

Quantifying the Activity of cis-Regulatory Elements in the Mouse Retina by Explant Electroporation

This protocol describes a simple and inexpensive way to quantify the activity of cis-regulatory elements (i.e., enhancer/promoters) in living mouse retinas via explant electroporation. DNA preparation, retinal dissection, electroporation, retinal explant culture, and post-fixation analysis and quantification are described.

Transcription factors within cellular gene networks control the spatiotemporal pattern and levels of expression of their target genes by binding to ...

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses specifically on antisense oligonucleotides (ASOs), though the techniques shown in the video here can also be used to deliver a plethora of other drugs to the CNS. Antisense oligonucleotides (ASOs) have the capability to knockdown sequence-specific targets 1 as well as shift isoform ratios of specific genes 2. To achieve widespread gene knockdown or splicing in the CNS of mice, the ASOs can be delivered into the brain using two separate routes of administration, both of which we demonstrate in the video.
The first uses Alzet osmotic pumps, connected to a catheter that is surgically implanted into the lateral ventricle. This allows the ASOs to be continuously infused into the CNS for a designated period of time. The second involves a single bolus injection of a high concentration of ASO into the right lateral ventricle. Both methods use the mouse cerebral ventricular system to deliver the ASO to the entire brain and spinal cord, though depending on the needs of the study, one method may be preferred over the other.

We describe a method to target drugs to the central nervous system by either implanting a catheter or performing a bolus injection into the right lateral ventricle in mice. We focus specifically on the delivery of antisense oligonucleotides. This technique is readily adaptable to other drugs and to rats.

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses ...

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution has been a long sought tool for neuroscientists in the systems-level search for the neural circuitry governing complex behavioral states. Optogenetics is a cutting-edge tool that is revolutionizing the field of neuroscience and represents one of the first systematic approaches to enable causal testing regarding the relation between neural signaling events and behavior. By combining optical and genetic approaches, neural signaling can be bi-directionally controlled through expression of light-sensitive ion channels (opsins) in mammalian cells. The current protocol describes delivery of specific wavelengths of light to opsin-expressing cells in deep brain structures of awake, freely-moving rodents for neural circuit modulation. Theoretical principles of light transmission as an experimental consideration are discussed in the context of performing in vivo optogenetic stimulation. The protocol details the design and construction of both simple and complex laser configurations and describes tethering strategies to permit simultaneous stimulation of multiple animals for high-throughput behavioral testing.

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Optogenetics has become a powerful tool for use in behavioral neuroscience experiments. This protocol offers a step-by-step guide to the design and set-up of laser systems, and provides a full protocol for carrying out multiple and simultaneous in vivo optogenetic stimulations compatible with most rodent behavioral testing paradigms.

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution ...

Simultaneous Electrophysiological Recording and Micro-injections of Inhibitory Agents in the Rodent Brain

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing system was developed that allows for the injection of a drug while recording electrophysiological signals from the affected neuronal population. This method provides a simple and economical alternative to commercial solutions. A glass pipette was modified by combining it with a hypodermic needle and a silver filament. The injectrode is attached to commercial microsyringe pump for drug delivery. This results in a technique that provides real-time pharmacodynamics feedback through multi-unit extracellular signals originating from the site of drug delivery. As a proof of concept, we recorded neuronal activity from the superior colliculus elicited by flashes of light in rats, concomitantly with delivery of drugs through the injectrode. The injectrode recording capacity permits the functional characterization of the injection site favoring precise control over the localization of drug delivery. Application of this method also extends far beyond what is demonstrated here, as the choice of chemical substance loaded into the injectrode is vast, including tracing markers for anatomic experiments.

Here, we craft a glass pipette with dual functions: inhibition of deep brain structures by microinjections of drugs and real-time monitoring of their effects through simultaneous electrophysiological recordings.

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing ...

Personalized Needles for Microinjections in the Rodent Brain

Microinjections have been used for a long time for the delivery of drugs or toxins within specific brain areas and, more recently, they have been used to deliver gene or cell therapy products. Unfortunately, current microinjection techniques use steel or glass needles that are suboptimal for multiple reasons: in particular, steel needles may cause tissue damage, and glass needles may bend when lowered deeply into the brain, missing the target region. In this article, we describe a protocol to prepare and use quartz needles that combine a number of useful features. These needles do not produce detectable tissue damage and, being very rigid, ensure reliable delivery in the desired brain region even when using deep coordinates. Moreover, it is possible to personalize the design of the needle by making multiple holes of the desired diameter. Multiple holes facilitate the injection of large amounts of solution within a larger area, whereas large holes facilitate the injection of cells. In addition, these quartz needles can be cleaned and re-used, such that the procedure becomes cost-effective.

We describe here a protocol for microinjection in the rodent brain that uses quartz needles. These needles do not produce detectable tissue damage and ensure reliable delivery even in deep regions. Moreover, they can be adapted to research needs by personalized designs and can be re-used.

Microinjections have been used for a long time for the delivery of drugs or toxins within specific brain areas and, more recently, they have been used to ...

Evaluation of Synapse Density in Hippocampal Rodent Brain Slices

In the brain, synapses are specialized junctions between neurons, determining the strength and spread of neuronal signaling. The number of synapses is tightly regulated during development and neuronal maturation. Importantly, deficits in synapse number can lead to cognitive dysfunction. Therefore, the evaluation of synapse number is an integral part of neurobiology. However, as synapses are small and highly compact in the intact brain, the assessment of absolute number is challenging. This protocol describes a method to easily identify and evaluate synapses in hippocampal rodent slices using immunofluorescence microscopy. It includes a three-step procedure to evaluate synapses in high-quality confocal microscopy images by analyzing the co-localization of pre- and postsynaptic proteins in hippocampal slices. It also explains how the analysis is performed and gives representative examples from both excitatory and inhibitory synapses. This protocol provides a solid foundation for the analysis of synapses and can be applied to any research investigating the structure and function of the brain.

A protocol for accurately identifying and analyzing synapses in hippocampal slices using immunofluorescence is outlined in this article.

In the brain, synapses are specialized junctions between neurons, determining the strength and spread of neuronal signaling. The number of synapses is ...

Recording and Modulation of Epileptiform Activity in Rodent Brain Slices Coupled to Microelectrode Arrays

Temporal lobe epilepsy (TLE) is the most common partial complex epileptic syndrome and the least responsive to medications. Deep brain stimulation (DBS) is a promising approach when pharmacological treatment fails or neurosurgery is not recommended. Acute brain slices coupled to microelectrode arrays (MEAs) represent a valuable tool to study neuronal network interactions and their modulation by electrical stimulation. As compared to conventional extracellular recording techniques, they provide the added advantages of a greater number of observation points and a known inter-electrode distance, which allow studying the propagation path and speed of electrophysiological signals. However, tissue oxygenation may be greatly impaired during MEA recording, requiring a high perfusion rate, which comes at the cost of decreased signal-to-noise ratio and higher oscillations in the experimental temperature. Electrical stimulation further stresses the brain tissue, making it difficult to pursue prolonged recording/stimulation epochs. Moreover, electrical modulation of brain slice activity needs to target specific structures/pathways within the brain slice, requiring that electrode mapping be easily and quickly performed live during the experiment. Here, we illustrate how to perform the recording and electrical modulation of 4-aminopyridine (4AP)-induced epileptiform activity in rodent brain slices using planar MEAs. We show that the brain tissue obtained from mice outperforms rat brain tissue and is thus better suited for MEA experiments. This protocol guarantees the generation and maintenance of a stable epileptiform pattern that faithfully reproduces the electrophysiological features observed with conventional field potential recording, persists for several hours, and outlasts sustained electrical stimulation for prolonged epochs. Tissue viability throughout the experiment is achieved thanks to the use of a small-volume custom recording chamber allowing for laminar flow and quick solution exchange even at low (1 mL/min) perfusion rates. Quick MEA mapping for real-time monitoring and selection of stimulating electrodes is performed by a custom graphic user interface (GUI).

We illustrate how to perform recording and electrical modulation of 4-aminopyridine-induced epileptiform activity in rodent brain slices using microelectrode arrays. A custom recording chamber maintains tissue viability throughout prolonged experimental sessions. Live electrode mapping and selection of stimulating pairs are performed by a custom graphical user interface.

Temporal lobe epilepsy (TLE) is the most common partial complex epileptic syndrome and the least responsive to medications. Deep brain stimulation (DBS) ...

Effects of Blast-induced Neurotrauma on Pressurized Rodent Middle Cerebral Arteries

Though there have been studies on the histopathological and behavioral effects of blast exposure, fewer have been dedicated to blast&#39;s cerebral vascular effects. Impact (i.e., non-blast) traumatic brain injury (TBI) is known to decrease pressure autoregulation in the cerebral vasculature in both humans and experimental animals. The hypothesis that blast-induced traumatic brain injury (bTBI), like impact TBI, results in impaired cerebral vascular reactivity was tested by measuring myogenic dilatory responses to reduced intravascular pressure in rodent middle cerebral arterial (MCA) segments from rats subjected to mild bTBI using an Advanced Blast Simulator (ABS) shock tube. Adult, male Sprague-Dawley rats were anesthetized, intubated, ventilated and prepared for Sham bTBI (identical manipulation and anesthesia except for blast injury) or mild bTBI. Rats were randomly assigned to receive Sham bTBI or mild bTBI followed by sacrifice 30 or 60 min post-injury. Immediately after bTBI, righting reflex (RR) suppression times were assessed, euthanasia at the time points post-injury was completed, the brain was harvested and the individual MCA segments were collected, mounted and pressurized. As the intraluminal pressure perfused through the arterial segments was reduced in 20 mmHg increments from 100 to 20 mmHg, MCA diameters were measured and recorded. With decreasing intraluminal pressure, MCA diameters steadily increased significantly above baseline in the Sham bTBI groups while MCA dilator responses were significantly reduced (p &#60; 0.05) in both bTBI groups as evidenced by the impaired, smaller MCA diameters recorded for the bTBI groups. In addition, RR suppression in the bTBI groups was significantly (p &#60; 0.05) higher than in the Sham bTBI groups. MCA&#39;s collected from the Sham bTBI groups exhibited typical vasodilatory properties to decreases in intraluminal pressure while MCA&#39;s collected following bTBI exhibited significantly impaired myogenic vasodilatory responses to reduced pressure that persisted for at least 60 min after bTBI.

Here, we present a protocol to describe methods for ex vivo vascular reactivity determination following a primary blast traumatic brain injury (bTBI) using isolated, pressurized, rodent middle cerebral arterial (MCA) segments. bTBI induction is accomplished using a shock tube, also known as an Advanced Blast Simulator (ABS) device.

Though there have been studies on the histopathological and behavioral effects of blast exposure, fewer have been dedicated to blast&#39;s cerebral ...

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 ...

Quantifying Subcellular Ubiquitin-proteasome Activity in the Rodent Brain

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Simultaneous fMRI and Electrophysiology in the Rodent Brain

Quantifying the Activity of cis-Regulatory Elements in the Mouse Retina by Explant Electroporation

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Simultaneous Electrophysiological Recording and Micro-injections of Inhibitory Agents in the Rodent Brain

Personalized Needles for Microinjections in the Rodent Brain

Evaluation of Synapse Density in Hippocampal Rodent Brain Slices

Recording and Modulation of Epileptiform Activity in Rodent Brain Slices Coupled to Microelectrode Arrays

Effects of Blast-induced Neurotrauma on Pressurized Rodent Middle Cerebral Arteries

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