Analyzing Supercomplexes of the Mitochondrial Electron Transport Chain with Native Electrophoresis, In-gel Assays, and Electroelution

All experiments were performed using hearts from C57BL/6N mice (wild type). Mice were anesthetized with CO₂ prior to cervical dislocation, and all procedures were performed in strict accordance with the Division of Laboratory Animal Medicine at the University of Rochester and in compliance with state law, federal statute, and NIH policy. The protocol was approved by the Institutional Animal Care and Use Committee of the University of Rochester (University Committee on Animal Resources).

1. CN and BN PAGE

NOTE: All equipment used for BN and CN PAGE must be free of detergent. To ensure this, wash all equipment with 0.1 M hydrochloric acid, followed by extensively rinsing with deionized H₂O.

1. Preparation
   1. Prepare anode buffer consisting of 25 mM imidazole, pH 7.0, at 4 °C; store at 4 °C.
   2. Prepare cathode buffer for CN or BN PAGE.
      1. For CN PAGE, use 7.5 mM imidazole and 50 mM tricine; add 0.5 g of sodium deoxycholate and 0.2 g of lauryl maltoside per liter of buffer. Adjust the pH to 7.0 at 4 °C and store at 4 °C.
      2. For BN PAGE, use 7.5 mM imidazole and 50 mM tricine and adjust the pH to 7.0 at 4 °C. Store at 4 °C.
      3. For light-blue BN cathode buffer, use 7.5 mM imidazole and 50 mM tricine. Adjust the pH to 7.0 at 4 °C and add 20 mg of Coomassie per liter of buffer. Store at 4 °C.
   3. Prepare extraction buffer (EB) with 50 mM NaCl, 50 mM imidazole, 2 mM aminocaproic acid, and 1 mM EDTA. Adjust the pH to 7.0 at 4 °C and store at 4 °C.
   4. Prepare 3x gel buffer (used for BN and CN gels) with 75 mM imidazole and 1.5 M aminocaproic acid. Adjust the pH to 7.0 at 4 °C and store at 4 °C.
   5. Prepare a loading buffer (LB) of 0.01 g of Ponceau S, 5 g of glycerol, and 5 mL of H₂O. Store at room temperature.
   6. Prepare Coomassie blue by adding 50 mg of Coomassie to 1 mL of 500 mM 6-aminohexanoic acid. Adjust the pH to 7.0 at 4 °C and store at 4 °C.
   7. Prepare 20 mM and 200 mM lauryl maltoside in H₂O. Make aliquots of 200 µL and store them frozen. Thaw the detergent before use.

Table 1: Quantities of Ingredients Needed to Pour 1 Mini- or Maxi-PAGE. The volumes used in this table are calculated for 1 mini- or 1 maxi-gel, 1.5 mm thick. The volume of AAB is based on a 40% stock solution. Light and heavy refer to the concentration of AAB. APS and TEMED are added after each column of the gradient mixer is filled with AAB solution.

2. Pouring and running gels
   NOTE: Use 3-8% or 4-10% acrylamide/bisacrylamide (AAB) gradient for CN or BN gels, respectively. Table 1 summarizes the quantities of buffer, AAB, H₂O, glycerol, ammoniumpersulfate (APS) and tetramethylethylenediamine (TEMED) used for a mini-gel (85 mm wide x 73 mm high x 1.5 mm thick) or maxi-gel (160 mm wide x 200 mm high x 1.5 mm thick). Assemblies of glass plates with CN or BN gels can be refrigerated in a bag with a few mL of 1x gel buffer or wrapped in paper towel wetted with 1x gel buffer for storage. The gels are stable for use for up to a week.
   1. To pour the gel, place the gradient mixer on an elevated stirring plate to ensure that the gel will flow by gravity into the prepared gel chamber.
   2. Fill the outflow chamber of the gradient mixer with 4.77/25 mL (mini/maxi) of the heavy solution (with a higher concentration of AAB).
   3. Gently open the stop-cock connection between the heavy and light chamber and allow a drop of solution to go through to the other side.
      NOTE: This pushes air bubbles from the connecting tube and stop-cock, which will prevent flow between the two chambers. This cannot be done if both sides have already been filled, because the equal pressure will prevent the bubble from moving through.
   4. Fill the other chamber of the gradient mixer with 4.72/25 mL (mini/maxi) of the light solution.
   5. Place a stir bar in the outflow chamber with the heavy solution and begin stirring. Use a stir bar speed that does not cause bubbling.
   6. Quickly add APS and TEMED to each chamber to initiate polymerization.
   7. Open the connection between the two chambers of the gradient mixer and allow mixing for a few seconds before opening the outflow chamber to pour the gel.
      NOTE: Gravity will drain both chambers equally, and mixing of the light into the heavy solution will slowly decrease the acrylamide density from the bottom to the top of the gel. Use the entire content that is in the gradient mixer to pour the gel.
      1. At the end, carefully mount the comb, with the wells inside the gel to avoid bubbles and layer mixing.
   8. Immediately wash the gradient mixer with ethanol to rinse out any gel. Rinse with water and pour the second gel. Let the gels polymerize (usually less than 20 min is needed for mini-gels).
   9. To run the gels, mount them into the electrode assembly clamp and fill the center/upper chamber with CN or light-blue BN cathode buffer. Wait a few minutes to check for leaks before adding anode buffer to the outer/lower chamber.
      1. Gently pull out the well combs and wash the wells with cathode buffer using a syringe or pipette.
      2. Run the gels in a cold room (4 °C) or completely packed in ice.
         1. For CN mini-PAGE, use 100 V for the first hour and 200 V until finished, usually an additional 1-1.5 h. Alternatively, run the CN mini-PAGE at 30-40 V overnight.
            NOTE: The focus here is on high-molecular-weight protein complexes, so protein complexes with a molecular weight of less than 140 kDa will run out of the gel. Shorter running times for electrophoresis can be used to retain low-molecular-weight complexes.
         2. For BN maxi-PAGE, use 100 V and run the gels overnight (about 18 h).
            NOTE: The current will be very low (< 15 mA), so a power supply that can handle these conditions is needed. At this point, the gels can be used for IGA or immunoblotting. In some cases, bands or lanes can be cut out of the gels using a razor blade on a glass plate for electroelution.
3. Sample preparation
   NOTE: Membrane-bound mitochondrial supercomplexes must be extracted from the inner mitochondrial membrane. To preserve mitochondrial supercomplexes, use either freshly isolated mitochondria or samples that have been frozen and thawed only once. The calculations/volumes below are given for mini-gels (the well of a 10-well comb in a gel 1.5 mm thick holds up to 35-40 µL) and maxi-gels (the well of a 15-well comb holds up to 200 µL). In addition, save an aliquot of each sample (usually 10 µL), to be run on a denaturing SDS gel for the detection of the voltage-dependent anion channel (VDAC), as a loading control.
   1. Place an appropriate amount (e.g., 10-50 µg of protein for mini-gels and 50 – 200 µg for maxi-gels) of isolated mitochondria or tissue homogenate in microtubes and centrifuge at 17,000 x g for 10-15 min at 4 °C.
      NOTE: This step removes some of the soluble mitochondrial matrix and/or cytosolic proteins.
   2. Aspirate and discard the supernatant and add extraction buffer to the amount desired to load onto the gel. Gently resuspend the sediment on ice. If desired, add a general protease inhibitor mix at this point.
      NOTE: Based on the equipment used here, 30 µL was used for a mini-gel and 100 µL for a maxi-gel, limiting the protein/buffer ratio to not more than 2 µg of protein to 1 µL of buffer.
   3. Add detergent (e.g., 2 µg of lauryl maltoside/1 µg of protein; see the Representative Results and Discussion for more information).
      NOTE: Generally, lauryl maltoside is used, but digitonin may also be used.
   4. Incubate on ice for 20 min. Gently mix at the beginning and occasionally during incubation by triturating and/or agitating the tube.
   5. Centrifuge at 17,000 x g for 10 min at 4 °C to remove any membrane and tissue fragments.
   6. Transfer the supernatant to a new tube. For CN samples, add 1 µL of LB for every 10 µL of sample volume; the total volume of the sample should be approximately 40 µL for a mini-gel and 130 µL for a maxi-gel. For BN samples, add Coomassie to the samples so that the ratio of dye to detergent is 1:4 (w/w).
   7. Load 30 and 120 µL of the samples into the wells of the mini- or maxi-gel, respectively. Use the remaining 10 µL from each sample for the denaturing SDS gel to detect VDAC as a loading control.
   8. To prepare the molecular weight markers, dissolve 1 vial of a high-molecular-weight calibration mix (see the Table of Materials for more information) in 60 µL of BN/CN gel buffer and add 120 µL of H₂0 and 20 µL of LB. Load 15 µL per lane.
   9. Run the gels as outlined in step 1.2.9.2.

2. In-gel Assays for Cx-I and Cx-V

NOTE: The assays are performed at room temperature. Take photos, scans, or images of the developing bands for documentation. (Important) Proteins cannot not be transferred onto nitrocellulose membranes after completing an IGA.

1. Cx-I assay
   1. Preparation.
      1. Prepare an assay buffer of 5 mM Tris in H₂O, with a pH of 7.4; store at room temperature.
      2. Dissolve 10 mg of NADH in 1 mL of assay buffer. Store in aliquots of 100 µL at -20 °C until use; avoid freezing and thawing.
      3. Weigh 25 mg of Nitroblue tetrazolium into a microtube.
      4. Prepare the fixative by diluting 5 mL of acetic acid in 95 mL of H₂O; store at room temperature.
   2. Performing the assay.
      1. Combine 10 mL of assay buffer with 25 mg of Nitroblue tetrazolium (final concentration: 2.5 mg/mL) and 100 µL of 10 mg/mL NADH (0.1 mg/mL). Add this to the entire gel, a lane, or an area of interest excised from a CN gel.
         NOTE: This can be done in a clear plastic or glass container. Please note that this assay cannot be performed after BN PAGE.
      2. Follow the development of blue bands after gentle agitation (rocker) for > 3 min.
      3. Fix the gel in 10 – 20 mL of acetic acid solution or wash in 5 mM Tris, pH 7.4 to stop the reaction. Take photographs for documentation (see the Table of Materials).
2. Cx-V assay
   NOTE: The Cx-V assay should be done using duplicate CN or BN gels, where one is incubated with 5 µg/mL oligomycin (a Cx-V inhibitor) to demonstrate Cx-V-independent activity.
   1. Preparation.
      1. Prepare an assay buffer with 35 mM Tris and 270 mM glycine; adjust the pH to 8.3 at room temperature. Store the buffer frozen in 50-mL aliquots, but check the pH after freezing and thawing.
      2. Prepare 1 M MgSO₄ in H₂O; store at 4 °C until use.
      3. Weigh 27.28 mg of Pb(NO₃)₂ into a microtube.
      4. Weigh 60 mg of ATP into a microtube.
      5. Dissolve 1 mg of oligomycin in 1 mL of ethanol. Store at -20 °C until use.
      6. Prepare a fixative by mixing 50 mL of methanol with 50 mL of H₂O; store at room temperature.
   2. Performing the assay.
      1. Incubate a gel, a lane, or an area of interest from a BN or CN gel for 2 h with gentle agitation (rocker) in 10-20 mL of assay buffer ± 5 µg/mL oligomycin (50-100 µL of 1 mg/mL in ethanol) at room temperature.
      2. After incubation, replace the buffer with 14 mL of fresh assay buffer and add, in order, 190 µL of 1 M MgSO₄ (14 mM), 27.28 mg of Pb(NO₃)₂ (5 mM), 60 mg of ATP (8 mM), and 75 µL of oligomycin (where necessary).
      3. Incubate with gentle agitation (rocker) and watch for a white precipitate, as bands on oligomycin-treated gels will give non-Cx-V-dependent bands.
         NOTE: The appearance of the precipitate may take several hours.
      4. Fix the gel in methanol-based fixative (e.g., 50% methanol), because acidic solutions will dissolve the lead precipitate. Photograph the results.
         NOTE: The lead precipitate does not interfere with the Coomassie staining of the gels.

3. Protein Transfer to Nitrocellulose or Polyvinylidene Difluoride (PVDF) Membranes

1. Prepare a transfer buffer of 25 mM Tris and 200 mM glycine. Adjust the pH to 8.3 and add 0.0005 g/L SDS and 200 mL/L methanol. Store at room temperature.
   1. Cut a nitrocellulose or PVDF membrane (pore size: 0.45 µm) with a razor blade or scissors to a size slightly larger than that of the gel.
      NOTE: Nitrocellulose membranes allow for Ponceau S staining to assess protein loading, while PVDF membranes yield more sharply defined bands.
2. Protocol.
   1. Soak the nitrocellulose membrane, the filter paper, and the sponges for at least 10 min in transfer buffer. Place the PVDF membrane in 100% methanol for 15 s or according to the recommendation of the manufacturer before placing it in transfer buffer. Place the open cassette of the transfer kit into a flat bowl with transfer buffer.
   2. Place the sponge and 1-2 layers of filter paper on the back side of the cassette. Remove the bubbles.
   3. Place the gel on the filter paper. Indicate the orientation of the gel by clipping a corner of the gel and/or the membrane.
   4. Place the membrane on top of the gel. Remove all bubbles.
   5. Place filter paper and a sponge on top of the membrane. Remove all bubbles in this sandwich.
   6. Close the cassette and place it into the cassette holder of the transfer kit.
   7. Transfer the proteins at 25 V for about 12-18 h.

4. Immunoblotting

1. Preparation.
   1. Prepare a Ponceau Stain (500 mL) by adding 25 mL of acetic acid and 0.5 g of Ponceau S to 475 mL of H₂O; store at room temperature (can be reused).
   2. Prepare Tris-buffered saline (TBS) with 200 mM NaCl, 25 mM Tris-Base, and 2.7 mM KCl. Adjust the pH to 8.0 and store at room temperature.
   3. Prepare TBS-tween (TBST) by adding 0.5 mL/L Tween 20 to TBS; store at room temperature.
   4. Prepare milk solids/TBST by dissolving 5 g of milk solids in 100 mL of TBST. Store at 4 °C and use within 3 days.
   5. Prepare BSA/TBST by dissolving 3 g of bovine serum albumin (BSA, fraction V) in 100 mL of TBST. Store at 4 °C and use within 3 days.
2. Protocol.
   1. When the transfer is finished, place the membrane into Ponceau S solution to visualize all transferred proteins. Label the position of the markers on the membrane with a pencil and document the Ponceau-S-stained membrane by photograph or scan.
   2. Wash the membrane 3 times for 10 min each with TBS under gentle agitation.
   3. Block the membrane with milk solids/TBST for 1-2 h at room temperature or overnight in a cold room with gentle agitation.
   4. Wash the membrane for 10 min with TBST under gentle agitation.
   5. Incubate with primary antibody overnight in the cold room under gentle agitation. Dilute the antibody (e.g., 1:1,000 for anti-ATP5A and -NDUFB6) in BSA/TBST.
      NOTE: Most antibodies give a better signal on membranes from native gels when incubated overnight.
   6. Wash the membrane for 10 min with TBST under gentle agitation.
   7. Incubate with secondary antibody for at least 60 min at room temperature uner gentle agitation. Dilute the antibody (1:5,000 to 1: 50,000) in milk solids/TBST.
   8. Wash the membrane 3 times for 10 min at room temperature with TBST uner gentle agitation.
   9. During the last wash, prepare the enhanced chemoluminescence (ECL) substrate according to the instructions of the manufacturer.
   10. Incubate the membrane with ECL substrate using instructions provided by the manufacturer.
   11. Detect the signal on the film using instructions provided by the manufacturer of the ECL substrate.

5. Electroelution

1. Preparation.
   1. Prepare an elution buffer of 25 mM tricine, 3.75 mM imidazole (pH 7.0 at 4 °C), and 5 mM 6-aminohexanoic acid.
      NOTE: Wear gloves at all times while handling the electroeluter and membrane caps to prevent contamination with external proteins.
2. On the day before using the electroeluter for native electroelution, perform the following:
   1. Soak the membrane caps (cut-off: 3.5 kDa) in elution buffer for 1 h at 60 °C. Transfer the caps to fresh elution buffer and soak them for 12 h or longer in the refrigerator.
      NOTE: A cut-off molecular weight of 3.5 kDa prevents the loss of small proteins that may easily dissociate from the protein complex of interest.
   2. Wash the electroeluter module, glass tubes, tank, and lid thoroughly with ethanol. Rinse with water and let the equipment dry.
3. On the day of native electroelution, do the following.
   1. Place a frit in the bottom (frosted) of each glass tube to be used. If necessary, place the glass tube in elution buffer and push the frit from the inside to the bottom of the tube.
   2. Push the glass tube with the frit into the module of the electroeluter. Wet the grommet with elution buffer and slide the glass tube into place. Ensure that the tops of the glass tubes are even with the grommet.
   3. Close the empty grommets with stoppers.
   4. Place a wet membrane cap at the bottom of the silicone adapter and fill the adapter with elution buffer. Slowly pipette the buffer in the adapter up and down to remove any air bubbles around the dialysis membrane.
   5. Slide the buffer-filled adapter to the bottom of the glass tube. Remove all bubbles that appear on the frit inside the glass tube.
   6. Fill each glass tube with elution buffer.
   7. Place the excised bands of the BN PAGE into the glass tubes (see step 1.2.9.3). Cut large pieces into smaller pieces. Ensure that the fill height within the glass tube is around 1 cm.
   8. Place the entire module into the tank.
   9. Add about 600 mL of cold elution buffer to the tank. Ensure that the silicon adaptor caps are in the buffer to prevent bubbles at the dialysis membrane.
   10. Place a stir bar at the bottom of the tank.
       NOTE: Stirring will prevent bubbles from sticking to the bottom of the dialysis membrane.
   11. Elute the proteins for 4 h at 350 V in a cold room.
   12. After the elution is completed, remove the electroeluter module from the buffer tank and place it into a sink or bowl.
   13. If a stopper was used, remove it to drain the upper buffer chamber. Otherwise, use a large pipette to remove the buffer.
   14. Remove the buffer from each glass tube and discard. Make sure that the silicone adapter stays in place and that the liquid below the frit is not disturbed or shaken.
   15. Carefully remove the silicone adapter, together with the membrane cap from the bottom of the glass tube. Pipette the content (about 400 µL) of the silicone cap into a microtube. With another 200 µL of elution buffer, rinse the silicon cap and add to the microtube. Repeat for all glass tubes.
   16. As the membrane caps may be reused, preserve the membrane caps by placing them into elution buffer containing 0.5 mg/mL sodium azide. Refrigerate them.
       NOTE: The eluate has low protein concentration and can be concentrated using a centrifugal filter devices.