DIGE 2D-PAGE


1. Determine protein concentration. 


2. Re-hydrate protein in Urea buffer so concentration is 1ug/ul, for a slightly harsher lysis Thiourea may be used. (Once lysis buffer is defrosted once it should not be refrozen and used again)

4. Vortex until protein completely dissolved.

5. Test the protein extract on pH strip. If not above pH 8.5, add 0.2 ul 0.5M NaOH. Flick, spin, test again.

6. Spin at 13,000 x g, RT, in microfuge, 10 minutes.

7. Pipet 50 ug of appropriate protein into tubes Cy3 & Cy 5.

8. Add 1 ul of appropriate dyes SEPARATELY to tubes Cy3 & Cy5; vortex briefly; spin briefly to collect liquid.

9. React 60 minutes on ice, in the dark.

10. Add 1 ul 10 mM lysine per 1ul CyDye to each reaction tube; vortex briefly; spin to collect liquid.

11. React 10 minutes on ice, in the dark.

12. Perform the Cy2 labeling. This labeling is done in the same fashion that you are labeling but at a larger scale (100 ug). The sample (pooled internal standard) is composed of Control and Experimental proteins from each of you. This pooled sample is labeled with Cy2 and when included in the gels, provides the analytical software with an internal reference to permit the normalization.

13. Combine your Cy3 & Cy5-labeled proteins in one tube. Add 100ug of the Cy2-labeled sample (standard) and mix with your Cy3 & Cy5-labeled samples. Total protein is now 150 ug.

14. Add 5ul of IPG buffer, pH3-10 or pH3-7 depending on the range you want.

15. Add trace amount of Bromophenol blue, Destreak Solution up to 500ul and vortex.

16. Pipet sample into focusing tray, aiming for the middle of the lane. Make sure to get it all over the bottom. Try to avoid bubbles.

17. Remove protective cover from appropriate IPG strip using tweezers and place strip gel side down into the rehydration well very carefully with the “+” or positive end of the strip aligned with the “+” or positive end of the tray. Ensure that the length of the gel is covered in sample. Raising and lowering the strip several times can help to spread the liquid, and all bubbles under the strip should be avoided or removed.

18. Cover the strip with mineral oil to prevent them from drying out.

19. Rehydration and Focusing:
                            a. Rehydration should be active for at least 12 hours with an active 50v.
                            b. Suggested method for 24cm focusing:
                                                    24cm IPG Strip
                                                    50v for 12 hours
                                                    500v for 1 hour
                                                    1000v for 1 hour
                                                    8000v for 9 hours
                                                    100v for 5hours
                                                    Minimum Needed: 67000volt-hours
                            c. The last step is a removal window step. This is used to ensure that after the method is completed focusing is maintained if    immediate removal does not occur.

20. Remove IPG strip from focusing tray, place gel side up on paper towel to remove oil from back of strip.

21. Wipe the front of the IPG strip off very carefully. Following focusing, strips can be frozen at –80 ºC if necessary due to lack of time. When ready to continue: unfreeze and continue with the re-equilibration steps.

22. Re-equilibrate strips for second dimension in re-equilibration tray, which require approximately 2mL per lane to cover a strip. Strips should be placed on gel as quickly as possible to prevent diffusion.
                            a. Prepare re-equilibration solutions.
                                                      Solution 1 10mL Stock Re-equilibration Buffer + 50mg DTT
                                                      Solution 2 10mL Stock Re-equilibration Buffer + 450mg iodacetamide
                           b. Cover strips with solution 1, gel side up, and soak strips on shaker for 20 minutes.
                           c. Decant solution 1.
                           d. Cover strips with solution 2, gel side up, and soak strips on shaker for 20 minutes.
                           e. Decant solution 2.
                           f. Cover strips with 1x running buffer, gel side up, and soak strips on shaker for 5 minutes.

23. Prepare agarose sealing solution: 0.5% agarose in running buffer plus a trace of Bromophenol blue. Heat up in a microwave to make sure agarose is dissolved.

24. Ready 2nd Dimension Ettan DALT tank:
                           a. Fill the tank with running buffer and connect tank tubes to a water cooler.
                           b. Let tank run for a while before beginning.

25. Get out pre-made 24cm gels. Click here for gel preparation recipe.

26. Rinse tops with ddH2O, make sure gel has no excess water on top of it before placing strip in.

27. Place strips into empty gel well and push down till it is flat against the bottom of the well and there are no bubbles trapped underneath it. An old comb cut in half is a good tool to use to push strips into place. Tweezers work just as well (be careful not to push too hard).

28. Pipette agarose sealing solution with a transfer pipet on to the top of strip to seal it into place. Allow time for agarose to solidify.

29. Place gel into tank. Gels should be placed evenly on each side of tank if able and they should all be facing the same direction. Use gel plate spacers to fill empty slots.

30. Pour more buffer until level is within fill mark. Put lid on tank and place voltage connectors in voltage box.

31. Start voltage box. For an overnight gel run you can set 1.5W per gel. 2.5W per gel may take 4-5 hours. 32. Run gels until the Bromophenol blue is off the end of the gel.

33. Take gels out, and scan for image.


Image taken from Amersham Biosciences.


Urea Lysis Buffer: 

8M Urea                                12.0 g
DTT                                        0.5 g
4% CHAPS                            1.0 g

Store in 1ml aliquots at -20ºC. 



Thiourea Lysis Buffer: 

7M Urea                                 3.8 g
2M Thiourea                          10.5 g
DTT                                        0.5 g
4% CHAPS                            1.0 g

Store in 1ml aliquots at -20ºC. 



Re-equilibration Buffer: 

Tris HCl (1.5M stock pH 8.8 w/HCI)     13.4ml     50mM
Urea                                                  144.14g    6M
Glycerol (87% stock)                         138mL      30%
SDS                                                      8.0g         2%
Bromophenol blue                                  Trace (add with pipette tip)

Store in 40ml aliquots at –20ºC. 



Running Buffer: 

Tris Base                            60.57g
Glycine                               288.27g
SDS                                    40.00g

Adjust volume to 2L. pH ~8.5. Dilute down to 1x running buffer before using to run your gels.


0.5% (w/v) Agarose Sealing Solution: 
1x SDS buffer                      100ml
Agarose Prep                       0.5g
Bromophenol blue               Trace (add with pipette tip)


10mM Lysine: 
L-Lysine                      0.018g
Store in 1ml aliquots at -20ºC.

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