This is more a problem for the lower bands. How hot gets your gel?

Desalting is a very good solution. Secondly re-suspend your sample in water rather than loading buffer. You are using low voltage. Your gels should not heat up at this voltage. I experienced that the gels run nicely at 120 V. Ensure that there should be no mixing of anode and cathode buffers while running the gels.

Do not shake your gels while staining and destaining process, as this might result in the migration of small peptides out of the gel. Reduce the amount of protein that you are loading. prepare the gel according to the attached protocol. Use colored marker (preferably Fermentas # SM1861), so that you see that the smallest peptide band doesn't escape the gel.

High salt is major problem in SDS-PAGE. In your case, protein marker is also not resolving properly. protein marker resolved only in one image, so in your samples i think there is no such problem. in gel preparation. In your gels. During gel preparation, seperating gel is not polymerizing uniformly. uniform polymerization of acrylamide play main role in protein seperation. if your gel is not properly polymerized then such type of problems occurs in SDS-PAGE.

I agree with Mahendra, something is wrong with the gel. Check the pH of all solutions (including the loading buffer) and get fresh acrylamide (it goes bad over time as a 40% stock and also as a dry powder-but slower). As acrylamide breaks down the pH decreases and this affects the way proteins will run. Also, during the running of the gel, acrylamide will break down, and this will be more rapid with old acrylamide stocks. Also check the age of your temed. How quickly does your gel polymerize? The gel should polymerize in 2-3 min. after it is poured (put some in a 1.5 mL test tube to time). Once the gel has polymerized, wait 1 hr before using to make sure it is completely polymerized. This is important for the smaller proteins/[peptides, that is where you gels have problems resolving bands.

Dear all...thanks for your suggestions. Please guide me what is good procedure for de-salting? I have all new stocks like APS, TEMED, Acrylamide. I always avoid mixing of cathode and anode buffer. Secondly can you tell me is their any method to desalt samples mixed with loading buffer?

Regards

Waqar

Wagar,

What buffer are your samples in and do they all have the same concentration of buffer/salt? I have run samples with 100-500 mM NaCl, just diluted them into running buffer and added 6X loading buffer and they all ran fine.

Hi waqar, I think you should use 10 % APS instead of 37.5%, because APS and TEMED catalysis the polymerization, since the concentration of the gel is 40% (really high) thefore the polymers at the base of the gel is always undere gravitational stress due to leakage of the gel at the bottom and a wave like pattern of polymer may be envisioned. In order to avoid this wavelike setting pattern i suggest that lower concentration of APS(10 %) will be useful.

You can use zip tips to desalt your samples

^I second this. Check out the protocol that Millipore provides to see how simple it is. Also, using fresh buffers is a must!

http://www.cbs.umn.edu/sites/default/files/public/downloads/ZIPTIP_PROTOCOL_msp2011.pdf

Dear All

thanks for your cooperation. I desalt the protein samples and also diluted them up to 4 times. Here is new pic that is bit improved. However, this time I got double band at same position (4kDa, most lowered band). Please suggest me what to do in this scenario

Regards

Your gel is much better than before. It could be possible that that there are two peptides of nearly same molecular weight, which are isomers. Cut out the band and send for N-terminal sequence and tryptic digest analysis, to know if there are two isomers.

Thanks Aisha for nice suggestion. I observed same bands in my othet batches.

I had similar problem, its usually because your not filling the buffer volume correctly in tank and not correctly fitting the lid.