I have one cationic peptide and I want to check it's stabilty in vitro by treating it with various proteases such as elastases, Trypsin etc. Want the protocol to do this by SDS PAGE. Thanks and Regards.
After incubating the peptide with the protease for the desired amount of time, you should add an inhibitor of the protease, such as PMSF, to quench the reaction. Then add concentrated SDS-PAGE sample buffer and proceed in the usual way. The reason for adding the inhibitor, instead of just relying on the sample buffer to quench the reaction, is that when you add the sample buffer it may make the peptide more susceptible to proteolysis by denaturing any structure it may have before completely inhibiting the enzyme.
Thanks Sir for this valuable info. Is there any specific amount of peptide and proteases to be added.... means any well used protocol for that. How much peptide should be added to how much of protease any reference paper if u can suggest.
The amount of peptide to use should be sufficient so that you can see the band on a gel with Coomassie Blue staining without having to concentrate the sample. The amount of protease will have to be determined experimentally by titrating the concentration in the reaction for a fixed length of time. The chosen amount should cause just minor degradation, so that if the protein becomes more sensitive to proteolysis you will be able to see more degradation.
I suggest that you first load different amounts of substrate peptide, stain the way you would stain for the assay, and quantify the band intensities. Plot intensity vs amount loaded. You will want to choose a substrate amount that his high but within the linear range. Then, you can either adjust the amount of enzyme that you use or the amount of time for incubating enzyme with the substrate, or both. The ideal is, by the end of the reaction, to be within the range of 20% to 80% of original substrate that is digested. It helps to set up several repetitions of the zero hour sample. This is very important because your calculations of the percent of substrate digested is very much dependent on the starting amount. How I usually prepare the zero hour control is to have the buffer and substrate, add the reagent you use to stop the reaction, then the enzyme. It also helps to use different time points for the reaction. As for stopping the reaction, since you are studying stability to proteolysis, it will be important for you to check to make sure that no changes occur between the time you stop the reaction and the time you run the gel. Also, if your final reaction mix is acidic, note the color of your sample after adding Laemmli sample buffer. It should be blue. If yellow, the SDS-PAGE will not run properly. You will need to add 1 or 2 microliters of 1 M NaOH to titrate the bromphenol blue back to its blue color before loading on to the gel.
Hi Amir,
Since you say peptide, it could be that is low molecular weight molecule (
Such a small peptide will be difficult to resolve by SDS-PAGE, even with a Tris-tricine gel. Reverse-phase HPLC, monitored at 214 nm, would be a better method.
I agree Dr. Shapiro, chromatogaphic approach followed by UV detection would be best for your peptide. Good luck!
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