Isolation of Mitochondria for Mitochondrial Supercomplex Analysis from Small Tissue and Cell Culture Samples

NOTE: The composition of all culture media and buffers is specified in Table 1 and details related to all materials and reagents used in this protocol are listed in the Table of Materials.

1. Mitochondria isolation from cell culture

NOTE: The minimum volume of cells assayed has been ~30-50 µL of packed cells (step 1.4). This can correspond approximately to at least two or three 100 mm cell culture plates or to one 150 mm plate at 80-90% of confluence, depending on the cell type (between 5 × 10⁶ and 10⁷ cells in L929-derived cells or MDA-MB-468). Efficient cell breakage is a critical step.

1. Grow adherent cells to approximately 80% confluence or suspended cells to an adequate density.
2. Harvest cells by centrifugation at 500 × g for 5 min (directly from the cell suspension in the case of non-adherent cells or after standard trypsinization with a trypsin-EDTA solution of 0.05% trypsin and 0.02% EDTA in 1x PBS, in the case of adherent cells).
3. Wash cells 2x with cold 1x PBS and sediment them by centrifugation at 500 × g for 5 min.
   NOTE: From this point, all the steps must be performed at 4 °C. Therefore, all the reagents must be cold and the tubes containing cells or mitochondria must be kept on ice.
4. After the second centrifugation, discard the supernatant, estimate the cell pellet volume (cpv), and freeze the cells at -80 °C for at least 15 min to facilitate membrane breaking in step 1.8.
   NOTE: The protocol can be stopped at this point and the cells can be stored at -80 °C for weeks. Usually, 10 or 15 mL graduated tubes with conical bottoms are appropriate.
5. Let the cells thaw slowly on ice.
6. Resuspend the packed cell pellet in a volume of hypotonic buffer equal to 7x the cpv (10 mM MOPS, 83 mM sucrose, pH 7.2) (e.g., 100 µL of packed cells in 700 µL of buffer).
   NOTE: The volume must be enough to obtain efficient pops (see NOTE at step 1.8).
7. Transfer the cell suspension into a Potter-Dounce homogenizer of the appropriate size and let the cells swell by incubating them in ice for 2 min. For example, for a volume of 700-800 µL of cell suspension, a 1 mL capacity homogenizer is adequate.
8. Break the cell membranes by performing eight to ten strokes in the homogenizer coupled to a motor-driven Teflon pestle rotating at 600 rpm.
   NOTE: When making the strokes, it is important to create a vacuum that will increase the cell breakage efficiency along with the hypotonic and freezing actions for swelling and fragilizing the membranes, respectively. If it is well done, a "pop" sound will be heard when pulling down the homogenizer with a quick movement at each stroke.
9. Add the same volume of hypertonic buffer to the cell suspension (7x the cpv) (30 mM MOPS, 250 mM sucrose, pH 7.2) to generate an isotonic environment.
10. Transfer the homogenate into a 10-15 mL tube and centrifuge in a fixed rotor at 1,000 × g for 5 min at 4 °C.
    NOTE: The original tube containing the unbroken cells can be used for this purpose.
11. Transfer the supernatant, which contains the mitochondrial fraction, into 1.5 mL polypropylene tubes.
12. OPTIONAL: To increase the yield of mitochondria, resuspend the pellet from step 1.10 in 7x the cell pellet volume of buffer A (10 mM Tris, 1 mM EDTA, 0.32 M sucrose, pH 7.4) by pipetting up and down and centrifuge at 1,000 × g for 5 min at 4 °C. Combine the new supernatant with the previous one before proceeding to step 1.13.
13. Centrifuge in a microfuge at 16,000 × g for 2 min at 4 °C to collect the mitochondrial crude fraction.
14. Discard the supernatant and resuspend each mitochondria-enriched pellet with 0.5 mL of buffer A combining the contents of two tubes into one and centrifuge under the same conditions as described in step 1.13. Repeat the same process until all the material is in only one tube.
15. Discard the supernatant, resuspend the final pellet with 300 µL of buffer A, and quantify mitochondrial protein concentration using the Bradford assay. Centrifuge again as before and proceed to section 3.

2. Isolation of mitochondria from small amounts of animal tissues

NOTE: The minimum amount of tissue to apply this protocol with a certain confidence depends on the cell type and its mitochondrial abundance but could be ~20-30 mg of tissue for most cases. The tissue samples can be fresh material or frozen samples. In the latter case, allow the samples to thaw in homogenization buffer placed on ice before starting the procedure.

1. Weigh or estimate as closely as possible the amount (mg) of tissue.
2. Cut the tissue with a pair of scissors and make 3-4 washes in homogenization buffer with the help of a strainer taking care to avoid losing the smaller pieces.
   NOTE: This step is more important in tissues like skeletal muscle or heart than in the brain or softer tissues whose cells can be easily disaggregated directly by the homogenization step.
3. Add fresh homogenization medium (4 mL per gram of liver, 10 mL per gram of heart, brown adipose tissue (BAT) or muscle, and 5 mL per gram of brain or kidney; see specific buffer composition for each case in Table 1).
4. Transfer the pieces of tissue with buffer to the homogenizer and follow the optimal homogenization and mitochondria isolation protocol depending on the selected tissue.
5. Isolation of mitochondria from liver, spleen, and kidney
   1. Homogenize with four to six up-and-down strokes in the Elvehjem-Potter with a motor-driven Teflon pestle at 600 rpm.
   2. Transfer the homogenized tissue to a sterile centrifuge tube. Centrifuge in a swinging rotor at 1,000 × g for 5 min at 4 °C.
   3. Fill 1.5 mL polypropylene tubes with the supernatant obtained in the previous step. Centrifuge for 2 min at 16,000 × g in a microfuge at 4 °C and proceed hereafter as described before (steps 1.13 to 1.15)
6. Isolation of mitochondria from heart and muscle samples
   1. Homogenize with six to eight strokes in the Elvehjem-Potter with a motor-driven Teflon pestle at 600 rpm. Transfer the homogenized tissue to a sterile centrifuge tube.
   2. Centrifuge in a swinging rotor at 1,000 × g for 5 min at 4 °C. Pour the supernatant into clean 1.5 mL polypropylene tubes.
      NOTE: A second homogenization of the pellet obtained in step 2.6.2 can be performed with 4-5 additional strokes and ½ volume of homogenization buffer to increase the mitochondrial yield. The new supernatant (after centrifugation again at 1,000 × g for 5 min at 4 °C) can be combined with the previous one before proceeding to step 2.6.3. An alternative to increase the crude mitochondria yield is to dissociate heart and muscle tissue samples in trypsin solution before homogenization^36,37. In this case, the trypsin has to be efficiently inactivated/removed before the mitochondrial membranes are solubilized with digitonin (step 3.2).
   3. Centrifuge at 16,000 × g in a microfuge for 2 min at 4 °C to obtain the crude mitochondrial fraction.
   4. Discard the supernatant and resuspend each mitochondria-enriched pellet with 0.5 mL of buffer AT combining the contents of two tubes into one and centrifuge under the same conditions as described in step 2.6.3. Repeat the same process until all the material is in only one tube.
   5. Discard the supernatant, resuspend the final pellet with 300 µL of buffer A (no BSA), and quantify mitochondrial protein concentration using the Bradford assay. Centrifuge again as before and proceed to section 3.
7. Isolation of mitochondria from brain
   1. Homogenize the pieces of the brain with 10-15 strokes using a Dounce-type glass homogenizer with a manually driven glass pestle. Transfer the homogenized tissue to a sterile centrifuge tube.
   2. Centrifuge in a swinging rotor at 1,000 × g for 5 min at 4 °C.
   3. Collect the supernatant into a clean centrifuge tube.
   4. Resuspend the pellet obtained in the previous centrifugation step in the same volume of medium AT used in the first homogenization. Re-homogenize the pellet by repeating the process described in step 2.7.1 using 5-10 passes and centrifuge the suspension in a swinging rotor at 1,000 × g for 5 min at 4 °C.
   5. Remove the supernatant and add it into the tube prepared in step 2.7.3. Centrifuge at 10,000 × g in a fixed angle rotor for 10 min at 4 °C to obtain the crude mitochondrial fraction.
   6. Discard the supernatant and resuspend the pellet into 1/2 the initial homogenization volume (step 2.7.1) of medium AT. Distribute the mitochondrial suspension into clean 1.5 mL polypropylene tubes and centrifuge at 16,000 × g in a microfuge for 2 min at 4 °C.
   7. Discard the supernatant and resuspend each mitochondria-enriched pellet with 0.5 mL of buffer AT combining the contents of two tubes into one and centrifuge under the same conditions as described in step 2.7.6. Repeat the same process until all the material is in only one tube.
   8. Discard the supernatant, resuspend the final pellet with 300 µL of buffer A (no BSA), and quantify mitochondrial protein concentration using the Bradford assay. Centrifuge again as before and proceed to section 3.
8. Isolation of mitochondria from BAT
   1. Homogenize with eight to ten up-and-down strokes in the Elvehjem-Potter with a motor-driven Teflon pestle at 600 rpm. Transfer the homogenized tissue to a sterile centrifuge tube.
   2. Centrifuge in a swinging rotor at 1,000 × g for 5 min at 4 °C.
   3. Fill 1.5 mL polypropylene tubes with the supernatant obtained in the previous step avoiding the upper fat layer.
      ​NOTE: A second homogenization of the pellet obtained in step 2.8.2 can be performed to increase the mitochondrial yield. The new supernatant (after centrifugation again at 1,000 × g for 5 min at 4 °C) can be combined with the previous one before proceeding to step 2.8.4.
   4. Centrifuge for 2 min at 16,000 × g for 2 min in a microfuge at 4 °C.
   5. Discard the supernatant and resuspend each mitochondria-enriched pellet with 0.5 mL of buffer AT2 combining the contents of two tubes into one and centrifuge under the same conditions as described in step 2.8.4. Repeat the same process until all the material is in only one tube.
   6. Discard the supernatant, resuspend the final pellet with 300 µL of buffer A (no BSA), and quantify mitochondrial protein concentration using the Bradford assay. Centrifuge again as before and proceed to section 3.

3. Preparation of samples for Blue Native analysis

1. Resuspend the mitochondrial fractions (obtained from mammalian cell cultures or tissues) in BN sample buffer (50 mM NaCl, 50 mM Imidazole, 5 mM Aminocaproic acid, 4 mM PMSF) to obtain a protein concentration of around 10 mg/mL. See Table 2 for expected yields for the different types of samples.
2. Solubilize mitochondrial membranes by adding digitonin (10% stock solution) to obtain a ratio of 4 g of digitonin/g of mitochondrial protein (4 µL of digitonin 10% stock for 10 µL of the mitochondrial suspension prepared in step 3.1).
   NOTE: The detergent-to-protein ratio (g/g) must be optimized for each type of sample to obtain reproducible results; 2-8 g of digitonin/g of mitochondrial protein was found to be optimal for most tissues^34,35.
3. Mix by gently pipetting up and down. Incubate samples on ice for 5 min.
   NOTE: After this step, the mitochondrial suspension should become clearer (shift from opaque to translucent); otherwise, it could indicate that the amount of detergent is insufficient for correct membrane solubilization.
4. Centrifuge at full speed in a microfuge (20,000 × g approx.) for 25 min at 4 °C to remove insoluble material.
5. Collect the supernatant in a fresh tube. Add to the supernatant a volume of 5% G-250 (Coomassie Blue G-250 5% in 0.75 M aminocaproic acid) equivalent to 1/3 of the initial resuspension volume (step 3.1, this would correspond to a final proportion of 1.6 g of Coomassie/g of protein) and mix by pipetting.
6. Keep on ice before loading on the gel or freeze aliquots at -80 °C.

4. Blue Native gel electrophoresis

NOTE: The electrophoresis is performed in the cold room (4-8 °C). If there is no cold room facility, a cooling block can be introduced in the electrophoresis tank. Commercial 3-13% native polyacrylamide gels²⁹ are used, but homemade gels of the desired gradient concentrations can also be used^38,39.

1. Load the upper and lower chambers with the cold cathode A and anode buffers, respectively. Alternatively, load the samples in the wells before carefully filling the upper chamber with cathode buffer A.
   NOTE: Commercial electrophoresis buffers are used in our laboratory, but they can be prepared as indicated in Table 1.
2. Load between 30 and 100 µg of mitochondrial protein.
   NOTE: Usually, 30-50 µg of protein (for WB), or 40-100 µg of protein (for IGA) are loaded per well in a 10 lane gel (0.5 x 0.15 cm wells). Cathode A buffer contains Coomassie Blue G-250 and has an intense blue color that makes it difficult to see the gel wells for sample loading. It is convenient to mark the center of each well, with a red marker, for example, to help in placing the pipette tip during this step. Native molecular weight markers can be useful when setting up the BN-PAGE technique to confirm that the gradient gels are properly formed and that the complexes and SCs pattern is the correct one.
3. Run at 80-100 V for 25-30 min and then at 160-180 V, limiting the current to 12 mA/gel, until the dye reaches the bottom of the gel (~125-165 min in total).
4. If in-gel activity assays (IGAs) are to be performed, substitute the cathode buffer A with cathode buffer B when the dye front is in the middle of the gel (~1 h after the beginning of the run).
   NOTE: Although the gels can be documented without any staining just after the run since some bands are visible (mainly complex V, which can be considered an "internal" maker with a MW of ~600 kDa) due to their binding to G-250 dye, usually, SCs and individual complexes will not be visible without staining or IGA assays. The gels to be used in IGA assays or for WB should not be stained or fixed.
5. Disassemble the gel from the cassette and continue with Coomassie blue staining (section 5), IGA analysis (section 6), or WB immunodetection (section 7).

5. Gel staining

1. Stain the gel for 10-15 min at room temperature (RT) using Coomassie blue dye solutions (Coomassie dye R-250 at 0.25% in 40% methanol, 10% acetic acid; stain for 10-15 min).
2. Destain with several washes (typically 4-5 x 15 min) in 40% methanol plus 10% acetic acid at RT.
3. Document the gel.

6. In gel-activity (IGA) assays

1. Prepare IGA solutions for the analysis of the different respiratory complexes before the end of the electrophoresis and maintain them in the dark (by using dark-colored plastic boxes or covering them with aluminum foil, for example). The composition of each buffer is detailed in Table 1.
2. Place the gel or the lanes to be used in a plastic box as small as possible to accommodate it.
3. Add enough volume of the appropriate solution to cover the gel (usually 5 or 10 mL for 5 or 10 gel lanes, respectively) and incubate at RT with gentle shaking (60-80 rpm) and away from light.
   NOTE: The time of incubation depends on the complex to be analyzed and on the nature and amount of sample loaded; CI and CIV are the easiest and fastest to give results. Thus, CI activity will usually start to become visible after a few minutes, while CII and CIV need ~30 min to be observed and CV ~1.5-2 h. The reaction can continue for hours in all cases and the rate of signal intensification can be reduced by moving the incubation to a cold room (4-6 °C), for example, for an overnight (ON) incubation. CIII activity works only for clear native electrophoresis (CN), not for BN³⁴.
4. When the appropriate bands have developed, stop the reactions by removing the assay solutions, washing 2x with distilled water, and fixing the gel with 40% methanol plus 10% acetic acid (except for CV which is fixed only with methanol) for 30 min.
   1. To analyze CIV activity after CI detection by IGA, wash the gel 2x with distilled water, document CI activity, incubate the gel (without fixing it!) with 50 mM potassium phosphate buffer (pH 7.2) 2x for 30 min, and then with the complete CIV reaction buffer until the complex IV bands appear (usually by incubating ON at RT or 4 °C).
5. Document the gels.
   NOTE: For ATPase activity, as the developed bands are white, when documenting the gel, place it on a dark background so that the clear bands would be visible.

7. Western blot analysis

1. Prepare transfer buffer according to Table 1 and keep at 4 °C until the end of the electrophoresis.
2. Place the gel in a tray and add transfer buffer. Incubate at RT for 10-15 min.
3. Cut a piece of PVDF membrane of the same size as the gel and activate it in methanol for 10 s under agitation. Wash several times with distilled water and add transfer buffer. Incubate at RT for 10-15 min with gentle shaking.
4. Prepare the transfer sandwich, from bottom to top, avoiding bubbles between the gel and the membrane, in the following order: black side of cassette-sponge-blotting paper-gel-membrane-blotting paper- sponge-clear side of cassette.
5. Close and lock the cassette and put the sandwich in the transfer tank in the correct orientation for transfer (black side towards the negative pole).
6. Fill the transfer tank with transfer buffer, add a magnetic stirrer to agitate the buffer, and connect the power supply at 80 V for 2 h or at 100 V for 1 h. Perform the transfer between 4 and 8 °C (for example in the cold room) and with a cooling block in the transfer system.
7. Retrieve the membrane and continue with a standard western blot protocol, using specific antibodies for the different respiratory complexes to be detected²⁹.