SDS-PAGE protocol: visualizing proteins on acrylamide gels

Peccoud Lab Protocol: visualizing proteins via sodium dodecyl sulfate-polyacrylamide electrophoresis (SDS-PAGE)

Introduction

Because cellular extracts contain thousands of different proteins at a wide range of concentrations, it is often difficult to detect and measure specific proteins in these mixes, even when proteins are expressed at high concentrations. The principles of protein gel electrophoresis are similar to those of DNA gel electrophoresis. Proteins are separated by size in a gel matrix by applying a current causing the molecules to migrate and sift through the gel towards the positively-charged anode. Larger proteins migrate slower through the gel. The main difference between protein and DNA gel electrophoresis are that not all proteins are negatively charged. Uniform negative charges are chemically added to the proteins – in our case we will be performing sodium dodecyl sulfate-polyacrylamide electrophoresis (SDS-PAGE), which uses the negatively charged detergent SDS to non-specifically bind to proteins and coat them. Proteins also tend to be more compact that DNA molecules, so the gel matrix must have smaller pores. Therefore, protein gels use polymerized acrylamide, which is also much tougher than agarose allowing gels to be much thinner and to undergo extensive handling during staining and blotting.

SDS-PAGE Materials

SDS-PAGE Method

  1. Wear gloves at all times.
  2. Measure out enough 2x sample buffer for all samples (i.e. a little more than the total sample volume) taking into account that you will need to add 1/10 the total volume of reducing agent (10x DTT).
  3. Pipet the desired volume (usually >40 uL) of protein sample (cell lysate/protein sample/cell pellet) into screw cap tubes.
  4. Add to each sample an equal volume of the reducing 2x sample buffer and mix by trituration and vortexing.
  5. Boil the samples for 5 min and centrifuge max speed for 1 min to pellet any cell debris.
  6. Remove Novex 16% Tris-Glycine precast gels from their packaging. Remove the strip of tape at the bottom and carefully lift out the comb in the top, slightly tilting the comb to one side to allow air to enter the wells.
  7. Place gels in the electrophoresis lower buffer chamber on opposite sides of the upper buffer chamber (the part with the electrodes). Ensure that the lower side of the gel tops (where the combs were) are facing each other towards the inside of the upper buffer chamber. The running buffer in the upper chamber will fill the wells of the gel.
  8. Place the gel tension wedge in the lower buffer chamber with the flat side of the wedge against the gel cassette and the lever out at the edge of the chamber. Ensure that the plastic of the gel cassettes is making uniform contact with the rubber gasket of the upper buffer chamber and that they are making contact with the bottom of the lower buffer chamber.
  9. Pull the lever of the tension wedge toward the gel cassettes. The lever should click into place without much force and the gel cassettes should be level with each other.
  10. Using a 100 mL graduated cylinder, carefully fill the upper buffer chamber between the gel cassettes with 1x Tris-Glycine running buffer (the SDS will generate too many bubbles to see the buffer level if poured too fast). Fill up to just below the upper edge of the cassette.
  11. Let the apparatus sit for 5 min to see if any buffer leaks into the lower chamber. If it does, loosen the lever, remove the wedge, upper buffer chamber, and gel cassettes, then pour the buffer into a beaker. Start again.
  12. If the buffer doesn’t leak into the lower chamber, fill the lower buffer chamber up to about the lower edge of the wells. Only about an inch of buffer is necessary to run the gels (it must cover the open slot of the cassette that was covered by the tape), but filling the chamber allows for better heat dissipation during the run.
  13. When pipetting samples into the wells place the tip vertically into the well close to the bottom and slowly dispense the sample, allowing it to drop to the well bottom.
  14. Carefully turn the chamber around to allow easier loading of the second gel.
  15. Carefully turn the chamber around to allow for easier loading of the second gel.
  16. Place the top on the Chamber (it only fits in one orientation)
  17. Insert the electrodes into the large PowerPac. The electrodes are too small for the PowerPac ports, so use some tape to keep the electrodes in place.
  18. Set up the PowerPac to run at 225 V for 30 min. After it stops, if the dye front is more than 2 cm from the bottom of the cassette continue running until it is about 1.5 cm from the bottom.
  19. When finished, turn off the PowerPac, remove the electrodes from the PowerPac,  remove the top of the chamber, loosen the lever and lift out the wedge, cassettes, and upper buffer chamber. Place the cassettes on paper towels, and pour the buffer into the “non-halogenated solvents” waste container. Rinse off the buffer chamber, wedge, and upper chamber with running tap water and place on the drying rack.
  20. With the gel cassette flat on the bench and the opening to the wells facing up, insert the gel knife between the two halves of a gel cassette and pry the plastic plates apart along their three sealed edges.
  21. Carefully lift the top plate at an angle stating at one of the bottom corners. The gel will remain attached to the bottom plate through the lower buffer opening.
  22. Using the gel knife, gently cut away the bits of gel between the wells and slide them off the plate onto the paper towel with the knife edge.
  23. OPTIONAL: If performing an immunoblot, it is often useful to determine how even the sample loading is by first running a gel for Bio-Safe staining (Coomassie blue). Loading of the samples can then be evened out for a second gel used for immunoblotting.

 

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