Hi Nancy,

two reasons for the problems with the 2D gels come to my mind:

1.) I think that you might have used too much protein for the first dimension isoelectric focussing. It can happen that this leads to aggregation/immobilisation of your proteins in your IEF strip. Use exactly the same amount you used for the silver-stained gel when you stain with Coomassie Blue.

2.) Also the staining protocol itself could be the problem. In fact, your first Coomassie gel does not look too bad (the second one is clearly overloaded). If you still have the first gel, you can try to silver-stain it. The result won't be too pretty but you could still see if there are more spots present in the gel at all. Be also aware that there are quite a few varieties of Commassie Blue (and staining protocols) around. Could you please give me some details on your staining protocol?

Best regards,

Christian

I don't think that your problem is with IEF, the spots seem to be resolved (see lower MW). I would persume that the depletion is not efficient. Could the abundant/overloaded protein in the third PAGE be albumine? 

I am using coommassie blue R-250. I wash the gel briefly with deionized water and then I incubate it overnight with the coommassie blue stain.

The second and third gel were obtained using the same sample. The fact is that the first time I obtained a gel with many spots in the upper side and then in  the others the resolution was lower and for some reason there are less spots. I have no idea what is going on. 

Unfortunately I don´t have the second gel to silver stain it.

Do you incubate your gels in fixation solution (most often these contain methanol and acetic acid) prior to Coomassie staining?

No I don't

I recommend to incubate your gels in fixation solution prior to staining with Coomassie:

I found a protocol (http://protocolsonline.com/protein-science/coomassie/) and modified it a little bit:

1.) Prefix gel in 50% methanol, 10% acetic acid, 40% H2O for 30 minutes to overnight.

2.) Stain gel in the above solution, with 0.25%-0.5% Coomassie Blue R-250, for 2 – 6 hours, until the gel is a uniform blue color. Staining is complete when the gel is no longer visible in the dye solution. Prior to complete staining, the gel will appear as a lighter area against the dark staining solution. This method will detect as little as 0.1µg/band.

3.) Destain for 4 – 24 hours in 5% MeOH, 7.5% HoAC, 87.5% H2O. Bands will begin to appear in 1 – 2 hours. Destain until background is clear (I sometimes do this overnight).

Good luck!

Regards, Christian

Well, I have run over 500 2-DEs and I find it very unlikely that this would significantly improve the spot pattern, proteins should be fixed by SDS in the PAGE and most Coomassie stain are made in MetOH/Acetic acid to preserve the fixation. I haven't even observed any significant loss in 2-DE stained with bio-safe Coomassie that requires removal of SDS prior the staining. Further, you wrote that there is a similar pattern in silver-stained PAGEs - these are fixed and with reasonable sensitivity even in basic protocols.    

If your three 2-DEs were made with the completely same sample, then I would consider protein stability (i.e. have you done several freezing-thawing cycles?). I have also obereved some loss in high MW when the IPG strips were not properly incubated in SDS prior the second dimension. In my protocol, there is 10 min reduction, 10 min alkylation and 5 min with buffer - i.e. 25 min with SDS. 

For more 2-DE troubleshooting, there is very nice manual by Westermeier&Naven - Proteomics in Practice (2002, Wiley) and you can also check Bio-Rad web. 

Good luck :-)

Martin mentions two important factors (protein stability and SDS incubation of IPG strips). I agree with him that you should check these first before attempting the pre-fixation of your gels...