|
|
|
|
|
|
|||||||||||||||||||||||||||||||||||||||||||
|
Separating gel |
10% |
|
|
Distilled water |
4.2 ml |
|
|
1.5M Tris-HCl<pH 8.8 |
2.5 ml |
|
|
10% SDS |
100 µl |
|
|
Acrylamide-bis (30% stock)* |
3.3 ml |
|
These materials may be mixed in the side-arm flask. Degas the solution by attaching it to a vacuum apparatus until bubbles stop forming at the surface (1-2 minutes).
Add:
|
10% ammonium persulfate |
50 µl |
|
|
TEMED |
5 µl |
Pour or pipette between the glass plates. Layer enough water over the surface of the acrylamide to form about a 2 mm layer.
2. When the separating gel has fully polymerized (20-40 minutes), carefully remove the water overlay by absorbing it with a piece of filter paper slid down between the two glass plates.
3. Allow any remaining acrylamide to polymerize in the flask and then scrap it out into a waste container in the hood. Rinse out the flask.
4. Prepare the stacking gel solution by combining the first four reagents in a 50 ml side-arm flask:
|
Stacking gel |
4.0% |
|
|
Distilled water |
6.1 ml |
|
|
0.5M Tris-HCl, pH 6.8 |
2.5 ml |
|
|
10% SDS |
100 µl |
|
|
Acrylamide-bis (30% stock)* |
1.3 ml |
|
5. Degas the solution for 2-3 min. or until there are no bubbles present using the vacuum pump.
6. Add 50 µl 10% ammonium persulfate and 10 µl TEMED to the flask and swirl. Be sure to mix thoroughly by gentle (NOT vigorous) swirling. Be careful about volumes because the wrong volumes of catalysts will not allow the gel to polymerize.
7. Immediately pour the stacking gel using a Pasteur pipette. Pour down the side of the gel apparatus near one of the spacers. Pour slowly and be careful to avoid the formation of bubbles. Place the comb at a slant between the two glass plates and lower gradually. This helps to push air bubbles out the top.
8. Allow the gel to polymerize for 20-40 min.
9. While the gel is polymerizing, prepare your hemolymph samples for loading. In a 0.5 ml microfuge tube add 1.5 µl of hemolymph, 6.5 µl water, and 2µl 5X SDS-PAGE sample dye. If a determination has been made of the protein concentration, use a volume of hemolymph containing 8 ug of protein plus 2 ul of 5X SDS-PAGE sample dye and use water to bring the final volume to 10 ul.
10. You will also be loading a molecular weight marker on each gel so prepare the marker. Pipette 10 µl of the marker solution into a 0.5 ml microfuge tube. Quick spin all the samples in the microfuge to mix the samples.
11. See step 14. Make the diluted running buffer.
12. Heat your samples and molecular weight markers at 100ūC in the water bath for 3 minutes just before loading. Do not store the samples on ice after heating.
13. After polymerization of the gel is complete, take the casting stand to the sink and rinse with water to remove any non-polymerized acrylamide. Then remove the comb by pulling it straight up, slowly and gently. Rinse the wells with distilled water at the sink using a water squirt bottle.
14. Remove the gel assembly from the casting stand and snap it into the cooling core (see instructions on the Mini-PROTEAN II Cell Assembly Guide at your table). Assemble so that two gels are in one unit.
15. Lower the cooling core into the lower buffer chamber. Dilute the running buffer from 5X to make 300 ml of 1X buffer. Add a small amount of 1X running buffer to the top chamber. Check for any leaks. If no leaks are found, pour ~115 ml of 1X running buffer into the top chamber.
16. Pour the remainder of the running buffer into the lower buffer chamber so at least the bottom 1cm of the gel is covered.
17. Briefly microfuge all of the sample tubes (again, be sure to balance!) for 4-5 sec. to collect the contents.
18. Load your entire sample into the well slowly and carefully using one of the special pipette tips provided by the faculty member. Hints:
19. Place the lid on top of the lower buffer chamber. Attach the electrical leads to the gel apparatus. Be sure to match red to red and black to black.
20. Attach the leads to the power supply and turn on power. Set the power at 200 volts; the current should be approximately 60 mA per gel.
20. Run the gel for approximately 45 min. or until the dye just begins to run out the bottom.
21. Turn the voltage to zero and then turn off the power supply. Disconnect the electrical leads.
22. Remove the cell lid and pull inner cooling core out of the lower chamber. Pour off the upper buffer into the sink and flush with running water.
23. Disassemble the units to obtain your glass plates holding the gel.
24. Push one of the spacers of the glass sandwich out to the side of the plates and twist to remove the top glass plate.
25. Carefully rip off the stacking gel and gently remove the separating gel from the second glass plate by grasping two corners and lifting off. These gels are very thin and may rip easily. Be careful! This is easier to do if you have wet fingers. Nick one edge of the separating gel for orientation. Record in your notebook which edge you marked.
26. Place the gel in enough Coomassie Blue stain to cover. Stain the gel for at least 30 min. More than one gel can be stained in the same container if there is a way to distinguish between the gels (such as a nick in the corner).
27. Clean all glass, spacers, combs, and the apparatus thoroughly with soap and water.
28. Transfer the gel from the stain to the destain solution. Cover the container with saran wrap. Pour the Coomassie Blue back into its original bottle using the funnel. Leave the gel in the destaining solution on the shaker for 30 min.-3 hr. with a sponge to help soak up the dye. Do not leave overnight. After destaining, transfer the gels to destain II. Pour the destain back into the "used destain" bottle for recycling or disposal.
29. Gels can be stored in a ziplock bag or they may be dried.
Click here for:
Hints for SDS-PAGE lab
Disclaimer: Supplier links are provided for your convenience only; these are not paid endorsements.
|
March 1999 |
