Quantifying Subcellular Ubiquitin-proteasome Activity in the Rodent Brain

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.

1. 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 °C freezer. Brains can be dissected the same day or at a later time.
2. 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).
   1. 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.
   2. 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.
3. 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 °C.

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.

1. Prepare the Lysis Buffer. Add 8.63 mL of ultrapure (deionized and distilled) water to a sterile 15 mL conical tube.
   1. To the conical tube, add 1,000 µL of 0.1 M HEPES, 15 µL of 1 M MgCl, 100 µL of 1 M DTT and 50 µL of 10% NP-40.
   2. Add 100 µL of protease inhibitor cocktail and 100 µL 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 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.
2. Prepare the Extraction Buffer. Add 5.925 mL of ultrapure (deionized and distilled) water to a sterile 15 mL conical tube.
   1. To the conical tube, add 2,000 µL of 0.1 M HEPES, 1250 µL of 50% glycerol, 15 µL of 1 M MgCl₂, 5 µL of 1 M DTT, 5 µL of 0.5 M EDTA and 600 µL of 5 M NaCl.
   2. Add 100 µL of protease inhibitor cocktail and 100 µL 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.
3. Remove the 1.5 mL centrifuge microtube containing one hemisphere of the ROI from the -80 °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.
4. Using a scalpel, transfer the frozen brain tissue to a 2 mL glass Teflon homogenizer. Add 500 µL of Lysis buffer to the Teflon tube.
   1. 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.
5. Using a 1,000 µL pipette, transfer the homogenized sample to a new 1.5 mL microcentrifuge tube. Place the tube on wet ice and incubate for 30 min.
6. Place tube in microcentrifuge and spin for 10 min at 845 x g and 4 °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 °C until Section 2.9.
7. Add 50 µL 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.
8. Place the tube in microcentrifuge and spin for 20 min at 21,456 x g, 4 °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.
9. Measure protein concentration of the Cytoplasmic and Nuclear extractions using the Dc protein assay (according to the manufacturer'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 °C until needed.

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

1. Prepare the TEVP buffer with 320 mM sucrose. Add 60 mL of ultrapure (deionized and distilled) water to a clean 100 mL beaker.
   1. To the beaker, add 1,300 µL of 1 M Tris (pH 7.5), 260 µL of 0.5 M EDTA, 100 µL of protease inhibitor cocktail, 100 µL of phosphatase inhibitor cocktail and 10.944 g of sucrose. Mix with a stir bar until sucrose is completely dissolved.
   2. Bring pH to ~7.4 by adding hydrochloric acid (HCl) dropwise to the solution using a pipette.
   3. 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.
2. Prepare the Homogenization Buffer. Add 60 mL of ultrapure (deionized and distilled) water to a clean 100 mL beaker.
   1. To the beaker, add 5,000 µL of 1 M Tris (pH 7.5), 3,000 µL of 5 M NaCl, 100 µL of protease inhibitor cocktail, 100 µL of phosphatase inhibitor cocktail and 2,000 µL 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.
   2. Bring pH to ~7.4 by adding HCl dropwise to the solution using a pipette.
   3. 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.
3. Remove the 1.5 mL centrifuge containing one hemisphere of the ROI from the -80 °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.
4. Using a scalpel, transfer the frozen brain tissue to a 2 mL glass Teflon homogenizer. Add 500 µL of TEVP buffer to the Teflon tube.
   1. 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.
5. Using a 1,000 µL 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 °C.
6. Collect the supernatant and transfer to a new 1.5 mL microcentrifuge tube using a 1,000 µL pipette. Centrifuge the sample at 10,000 x g for 10 min, 4 °C. The original pellet (P1) contains nuclei and the large debris and can be discarded.
7. Transfer the supernatant to a new 1.5 mL microcentrifuge tube. This is a Cytosolic Fraction. Add 50 µL of Homogenization buffer to the pellet (P2) and resuspend by pipetting until no solid material is visible.
8. Centrifuge the sample at 20,000 x g for 10 min, 4 °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.
9. Measure the protein concentration of the Synaptic fraction using the Dc protein assay (according to the manufacturer'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 °C until needed.

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 °C.

1. Plate reader settings: Pre-warm to 37 °C and hold through the run.
   1. 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.
   2. 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.
2. Reconstitute the 10x assay buffer provided in the kit with 13.5 mL of ultrapure water. Add 14 µL 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 °C until needed and is stable for several months.
3. Reconstitute the AMC standard provided in kit with 100 µL of DMSO. Perform this step in the dark or under low light conditions, as the standard is light sensitive.
4. Create a stand curve of AMC using the reconstituted standard, in the dark or under low light conditions.
   1. In separate 0.5 mL microcentrifuge tubes, add 16, 8, 6.4, 3.2, 1.6, 0.8, 0.4 and 0 µL of the AMC standard, which corresponds to 20, 10, 8, 4, 2, 1, 0.5 and 0 µM AMC concentration.
   2. To these tubes, in the same order, add 84, 92, 93.6, 96.8, 98.4, 99.2, 99.6 and 100 µL 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.
   3. Store all diluted standards on ice in the dark until needed.
5. Reconstitute the proteasome substrate (Suc-LLVY-AMC) provided in kit with 65 µL 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 µL per well. This equates to 220 µL need + 30 µL for pipetting errors, requiring 12.5 µL of substrate and 237.5 µL of 20S Assay buffer.
6. 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 µg is sufficient for any subcellular fraction.
7. Bring the sample well volume to 80 µL with ultrapure water. The amount added depends on the volume of sample added. For example, if sample 1 was 4.5 µL of protein and sample 2 was 8.7 µL, then the amount of water needed will be 75.5 µL and 71.3 µL, respectively. In two separate wells, add 80 µL 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.
8. Add 10 µL of 20S Assay buffer to each well, including Assay Blanks. A repeater/automated pipette is recommended here to ensure consistent assay volume across wells.
9. Optional: At this stage, introduce in vitro manipulations if desired; this will require that each sample has an additional 2 wells per treatment, including the vehicle. If so, add 5-10 µL 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 °C incubator for 30 min.
10. Turn off the lights or enter a dark room. Add all 100 µL of diluted AMC standards to a new well; each standard will have a single well.
11. In the dark, add 10 µL 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.
12. 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
13. At end of the kinetic run, export raw 360/460 fluorescent values to Microsoft Excel.
    1. 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.
    2. Take the highest AMC standard well average and divide by the known concentration (20 µM). 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.
    3. 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.
    4. Subtract the normalized Assay Blank value from each normalized sample value for all 5 scans.

5. Quantification of Linkage-specific Protein Ubiquitination

1. 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.
   1. For denaturing, mix the normalized samples with an equal volume of Laemmli sample buffer supplemented with β-mercaptoethanol to a percentage of 5% by volume, per the manufacturer's instruction.
   2. For quantification of all monoubiquitination and polyubiquitination modifications regardless of linkage, use a pan-ubiquitin antibody.
   3. To recognize all polyubiquitinated proteins, use an ubiquitin antibody that does not cross-react with monoubiquitination.
   4. For linkage-specific polyubiquitination, use antibodies that can detect Lysine-27, Lysine-48, Lysine-63 and linear (M1) polyubiquitination.
2. 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.
   1. 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).
   2. Incubate the membrane with the secondary antibody and redevelop in order to confirm that the membrane was stripped of primary and secondary antibodies properly.
   3. 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.
3. 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.
   1. 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.
   2. Subtract out the background, which is calculated as the mean optical density of the background immediately surrounding the column on all sides.