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No protein spots in 2D gel - Silver staining! - (Aug/13/2010 )

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Hello everyone, I've been having the following problem:

I run 15% SDS-PAGE gels, at about 100 mA for 2 hours. When I do only 1D my samples show up fine, lots of protein bands. However, when I use IPG strips (3-10), do IEF overnight and run the gels, no spots appear... The colored MW markers run normally in the gel, I can see them fine, but no spots appear. I initially thought that maybe my agarose sealant was the problem, since it didn't solidify very well, but I made a fresh batch and now it's fine.

Common problems I have read about no spots in silver staining include:

- temperature (too low, or too high), which is not the case,
- wrong silver nitrate dilution (also not the case, and it's a new reagent from Sigma)...
- Using bicarbonate instead of carbonate for developing solution (also not the case...)
- the only problem I could think of was the thiosulphate, although I have used the same batch before and it worked FINE... and when I used NEW GE healthcare thiosulphate no spots appeared either...

What bugs me the most is that I KNOW the MW markers are there, and not getting stained!!!

Anyone help? Suggestion?

Thanks a lot!


You should post the detail steps of your experiment.
How much are you loading your samples?
How long were the IPG strips soaked in Equilibration buffer (including 1st and 2nd)?


Hello, thank you for your reply... I loaded the limit according to the GE Healthcare Handbook, of 20 ug of protein per IPG strip. I also followed their 15 min rule for 1st and 15 min for 2nd equilibration, rinsed the strips a last time in 5 mL equilibration buffer rapidly and then sealed them with agarose and ran them at a constant rate.

I rehydrated them overnight, and did everything perfectly... is there any more information I can give you?

Thanks a lot... I'm hysterical trying to understand what may have gone wrong... :(


What was the length of the IPG strip you used?
I think you can increase the amount of loading samples. GE Healthcare Handbook just is a recommendation. The amount of loading is based on the type of sample (from cells, tissue, membrane proteins, nuclear proteins and so on). If your sample is complexity (including thousands of different proteins) you should increase your loading amount. Furthermore, the length and pH rang of the IPG strip is the factor you should consider. pH 3-10 maybe need a high sample amount.
Samples from tissue 250 ug for 24cm pH 3-10 IPG strip (silver strain)
180 ug for 18cm pH 3-10 IPG strip (silver strain)


Hello, are you sure about those amounts? pH 3-10 13 cm recommendation was 10-20 ug protein only!

Actually, I discovered my 2D quant-kit method leaves to be desired, and I think it is not very accurate, how do you quantify your proteins? Bradford, Lowry, kits?

I will try to up the amount of protein today, and let you know how it goes, but am still worried about my protein quantitation method...


I quantify proteins using 2D Quant-Kit (GE Healthcare). Bradford has a bad compatibility with Urea, Thiourea and DTT. So I recommend 2D Quant-Kit for quantification of proteins dissolved in Urea buffer.


actually, my proteins are biliary proteins diluted with milli-q water only... but u don't get much protein loss using that kit?


There is not any problem to using the Kit for protein quantification.
How do you prepare your sample?
do the proteins precipitated and dissolved in lysis buffer (Urea buffer)? If not, it will make bad IEF results, because of salt, fat, saccharide… in samples.


Hi, I use GE's sephadex columns G-25 with a exclusion size of 5 kDa to remove salts and other small interferents, diluting with water. And I don't solubilize them with and Urea, only with the solubilization buffer that GE recommends.. and no boiling either... But my GE 2D quant kit results give me extremely low protein values, and I think I may be getting some pretty big losses... but if u've never had a problema... I really don't know what else to do..!



The following method may be helpful to you.

Bile fluid samples were sonicated and centrifuged at 16 000 g for 15 minutes at 4 ℃ to remove debris, nucleic acid and mucins as a preliminary separation. For sample delipidation, each 1 ml of preliminary separated sample was then mixed with 250 µl of CleanasciteTM HC (Ligo-Chem, Inc., Fairfield, NJ, USA) followed by rotation for 1 hour at 4 ℃. After incubation, each sample was centrifuged at 16 000 g for 1 minute to clear away the formed lipid-micelles, and the supernatant was transferred to a new tube. In order to reduce the salt concentration and other contaminants, a commercially available microcentrifuge filtration device (YM-3, molecular mass cut-off at 3 kD; Millipore, Bedford, MA, USA) was used to wash away contaminating species.

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