My samples contain protein and likely green tea compounds. After my sample preparation I added NuPAGE reducing agent (Novex) and Novex NuPAGE sample buffer and heated the samples for 2 min at 85 degree Celsius (according to the manufacturer). I then stored my samples in the freezer. Samples then were blue from the sample buffer. I had to run my samples again weeks later and I noticed that first time I took them out 2 of my samples (was a digestive study..so had different aliquots) were not blue anymore but greenish/brown. A week later even now, they were all brownish in color. SDS-PAGE results look very similar, however on the latest PAGE I have smearing. Doesn't seem like I lost proteins but I wonder if the smearing comes from oxidation? pH changes? Anyone know why such frozen samples turn brown? I highly suspect reoxidation or change in pH...
Any suggestions are greatly appreciated.
Thanks!!
The blue dye (Coomasie blue) is also a pH indicator, so any change to the red (first greenish and then to brown-red) indicate an acidification of your sample.
I get some similar results with "home-made" sample buffer when it was prepared with out-of-date Tris-HCl buffer.
I suggest you check the best-date of your solutions, as could be one of them is "damaged" and the pH would not be the one described.
there's no solution for your treated samples, at least I don't know if you can 'correct' the pH of the samples.
The blue dye (Coomasie blue) is also a pH indicator, so any change to the red (first greenish and then to brown-red) indicate an acidification of your sample.
I get some similar results with "home-made" sample buffer when it was prepared with out-of-date Tris-HCl buffer.
I suggest you check the best-date of your solutions, as could be one of them is "damaged" and the pH would not be the one described.
there's no solution for your treated samples, at least I don't know if you can 'correct' the pH of the samples.
It would seem to me like some oxidation and pH change is occurring. To minimize this I suggest you try to run your samples immediately after prepping them (go from 85C denature step to ice for a couple of minutes before loading in a gel) instead of putting them in storage. Once you transfer the proteins from the gel to your membrane you can dry the membrane out and practically keep it forever. If you run 3 to 4 gels you can have lots of back up and time saved if a membrane goes wrong.
Thanks guys! Your thoughts are greatly appreciated. Interesting is that I have other such frozen/stored samples that contain a different kind of plant/fruit polyphenol set in the protein sample and those did not change color. Was just the green tea samples. But yes, I finally realized that the DTT in my reducing agent (bought commercially) is not stable, so you should not add samples buffer and reducing agent to your protein sample and freeze it. Also my solution was at an acidic pH when I froze them so I had likely pH effects as well as precipitation from plant compounds (that were invisible in solution when I froze the samples) and my proteins...
I am running a test SDS PAGE now with my original frozen stock solutions (in acidic solution no detergents etc.) to see if I had degradation over time/missing any protein bands..in any way I still deal with the protein/polyphenol precipitates.
I'll redo my experiment from scratch and then store my samples in SDS sample buffer right away until I need them and then add reducing agent and heat denature them. That should fix my pH issue as well as possible aggregations and oxidation processes due to unstable DTT. Hopefully :D
Josiah:
I didn't know you could blot your gel onto a PVDF membrane and practically dry and keep it as long as wanted for Western blots???! Interesting. How do you store the membranes until you need them? Will the blotted proteins not be affected in some way over time?
Best
Nat
I use nitrocellulose membranes, but I don't think there is a major difference between NC and PVDF other than cost and protein binding preference. For all my applications though NC membranes do the trick just fine.
The way I dry the membranes is by just doing a standard TBS-T wash procedure (3 washes for 5 min each) after the protein has been transferred to the membrane. Then pour off the wash buffer and let your membrane air dry for a few hours to overnight in a drawer. When it's completely dry you can seal it up in a ziploc bag and just keep it around. Depending on your application the proteins should be good indefinitely in a cool, dry, dark location like a drawer. And when using just give a quick wash in TBS-T and you're good to go.
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