Running SDS-PAGE Gels
To run an SDS-PAGE gel with the end goal of visualizing the protein on the gel
- Do you need to know the amount of protein (ug) to load, or does it not matter? If you need to know, you must run a Bradford on your protein samples
- Do you know if you have enough loading buffer, running buffer, and gel stain (Coomassie, silver, etc.)?
- Do you have a shaker that you can use to rock the gel back and forth? If not, there may be some shakers in Bio Superlab
- Are you planning to do a Western blot with this SDS-PAGE gel? This protocol is strictly for visualizing protein on the SDS-PAGE gel, which is not necessary if you want to run a Western blot. Also, doing a Western necessitates a few changes to the SDS-PAGE protocol, which are NOT outlined in this protocol for simplicity.
1. Set the hot block to 95 degrees C to give it time to heat up. Fill enough wells with water to hold all protein sample tubes
2. Set up the gel apparatus
- a. Remove gel from package and remove green tape at bottom
- b. Use a paper towel to dry the gel
- c. Gently push the gel cassette out using your thumbs. Make sure the comb comes out evenly.
- d. If running only one gel, grab a buffer dam (i.e., a “dummy” gel cassette)
- e. Place gel and buffer dam in BioRad tetra system, making sure they fit snugly inside without falling out (have to make sure the cassettes are wedged inside the two half-circles at the bottom ledge)
- f. The gels must be on opposite sides of each other, with the lane numbers facing outwards. In the case of the buffer dam, which has no lane numbers, there should be text that indicates how to orient it (“gasket” = inside of the tetra system)
- g. If two gels are run, just orient them with the lane numbers facing outwards
- h. Close using the green clamps.
- i. Place the tetra system in the gel box, with the black and red markers on the tetra system aligned with the black and red markers on the gel box
- j. Prepare a 1X solution of Tris/Glycine/SDS running buffer, invert several times to mix thoroughly, and pour into the tetra system itself (NOT the whole gel box) until you fill it to the top without spilling over
- i. How to make 1 L 1X solution: 100 mL of 10X running buffer in 900 mL of MilliQ water
- k. Lift the tetra system out of the gel box and check underneath for any buffer leakage
- l. If there is no leaking, pour running buffer in the gel box until you reach the “2 gel” line marked on the front of the gel box if you are running one (+buffer dam) or two gels. Fill to “4 gel” line if you are running three (+buffer dam) or four gels
- m. If there is leaking, then the gel cassette and/or buffer dam are likely oriented incorrectly. Fix accordingly.
3. Prepare 192 uL 5X Laemmli buffer + 8 uL BME in a 1.5 mL microcentrifuge tube. This is the loading buffer.
- a. Laemmli buffer is from Amy’s lab
- b. BME is a reducing agent that breaks disulfide bonds, should be in the acids section of MEE lab
4. If you use a gel with wells that can hold 50 uL, and you want to load maximum protein, prepare 40 uL protein sample + 10 uL loading buffer (1X) in 1.5 mL microcentrifuge tubes. Pipette up and down to mix.
- a. If you don’t need to load maximum protein, load 20 uL sample + 5 uL loading buffer instead. If you need the loading sample to reach a certain volume, you can either add more protein sample or molecular water.
- b. Negative control should also be prepared with loading buffer.
- c. Ideally, you should load equal amounts of protein in each well, but the only way to know the amount is to run a Bradford assay beforehand. Make sure you know whether or not you need to use the Bradford assay.
5. Heat the protein samples at 95°C in the water-filled wells of the hot block for 5 min to unfold the protein
6. Quick spin samples for 15 seconds to get all the condensation down into the rest of the sample
7. Pipette up and down to mix
8. Load 10 uL molecular weight marker (Precision Plus Protein™ Dual Xtra Prestained Protein Standards) into the first well, and load the protein samples into the rest of the wells
- a. Technique: Stick the tip against the front of the well, making sure you’re actually inside the well (there is a thin gap in which the tip must enter—if you’re using a P200 pipette tip, you should feel resistance). Move the tip downward halfway into the well, then dispense without going to the 2nd stop in order to prevent air bubbles. Remove the tip carefully without sucking up sample.
- b. It is recommended to load 10 uL of sample at a time (i.e., 5 x 10 uL per 50 uL sample) using a gel loading tip or P10 tip so that the tip is thin enough to easily fit in the wells. Using a P200 pipette tip to load the entire sample at once is quicker, but may lead to loading errors since it’s difficult inserting and removing the tip from the well.
9. Put on the electrodes/lid, making sure the cathode and anode on the lid match the ones on the tetra system
10. Run the gel for 1-1.5 hours at 75 V for good resolution, or 30-45 minutes at 150 V for worse resolution.
11. Stop the current when the dye front reaches the bottom of the gel
12. Remove the electrodes/lid, remove the tetra system, unclamp, and retrieve the gel cassette(s)
13. Crank open the gel using a spatula at the indicated arrows along the sides of the gel cassette (our 4-20% BioRad gels have these arrows, other gels may differ)
- a. Technique: start by inserting the spatula into the gap of the top-right arrow, lift up until that portion of the cassette breaks open a bit, then repeat with another gap further below, working your way toward the bottom-right arrow. Then, repeat starting with the top-left arrow, working your way toward the bottom-left arrow. The front cassette should come off if done correctly.
14. Slide the gel down into a small plastic container filled with tap water
15. Wash the gel in the water for 5 minutes, with agitation, and repeat 3 times (each time with fresh tap water)
16. Remove the water and add 50 mL Coomassie stain
17. Incubate the gel for 1 hour using agitation (rocker)
18. Rinse the gel with water for 30 minutes using agitation
19. Remove the water and image the gel in Superlab using the Coomassie dye setting and white plate