NuPAGE™ Tris-Acetate SDS Buffer Kit (for Tris-Acetate Gels), Contains 1 ea. LA0041, NP0004, NP0005, NP0007 - FAQs

DTT is not stable, so it must be added and the reduction performed just prior to loading your samples.

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Precipitation of the LDS or SDS at 4 degrees C is normal. Bring the buffer to room temperature and mix until the LDS/SDS goes into solution. If you do not want to wait for it to dissolve, you can store the sample buffer at room temperature.

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While they are both Bis-Tris based gels, the chemistries are very different since Bolt gels are optimized for western blotting. Another key difference is the wedge well design of the Bolt gels, which allows larger sample volumes to be loaded.

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No, CTAB will not work with any of our gels except for the NuPAGE Tris-Acetate gels. To use CTAB, you would need to use a running buffer of 50 mM acetic acid and 50 mM beta-alanine in equal concentrations. You would also need to switch the electrodes. Since CTAB is a cationic detergent, this would establish conditions for running a basic protein towards the anode (into the gel).

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One can use 0.002% thioglycolic acid in the upper buffer reservoir. This is a good scavenger of free radicals. The reference to this is described by Hunkapiller et al, Methods of Enzymology, (91), 399, 1983. Caution should be taken when using this method since this compound is both toxic and expensive. In addition, the TGA must be fresh as it tends to become oxidized itself over time. Oxidized TGA will actually promote sample re-oxidation.

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Copper or zinc staining is a rapid, sensitive method for detection of protein bands . ~10 ng reduced BSA on NuPAGE Bis-Tris gels can be detected with both the copper and zinc stain.

Copper Stain: Staining solution - 0.3 M CuCl2
After electrophoresis, remove the gel from the cassette and equilibrate the gel in 100 mL of 1X running buffer* for 15 minutes. Immerse the gel in 100 ml of 0.3 M CuCl2 solution for about 5 minutes (the protein band will appear as a negative stain with a blue background).

Zinc stain: Staining solution - 0.2 M Imidazole and 10 mM ZnCl2
After electrophoresis, remove the gel from the cassette and equilibrate the gel in 100 mL of 1X running buffer* for 15 minutes. Place the gel in 100 ml of 0.2M Imidazole solution for 10 minutes. Next immerse the gel in 100 ml of 10 mM ZnCl2 solution for about 5 minutes (the protein will appear as a transparent band with a white background).

*The 1X running buffer can be the buffer from the electrophoresis tank after run (MES, MOPS). However, for better contrast of the band, the 1X Tris-Glycine SDS running buffer is recommended.

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We do not recommend using gels past their expiration date because over time, the polyacrylamide hydrolyzes to acrylic acid and ammonia and this will affect the resolution of the proteins. Breakdown of polyacrylamide matrix is identified by:
- Ghost bands and doublets, seen first in the high molecular weight proteins
- Smiling of dye front across the gel, with bands in outer lanes becoming very slanted - proteins run slower there due to change in pH and pore size over time.


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This is not really recommended, but it is always possible to increase run time by lowering the voltage of the run. In general, the relationships are linear - i.e., decreasing voltage by half will double the run time.

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If you are running the gel at constant voltage, you do not need to increase the voltage regardless of the thickness of the gel.

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Using constant voltage allows the current and power to decrease during the run, providing a safety margin in case of a break in the system. Having lower power is a safety benefit and will also decrease the chances of experiencing an overheating of the gel. Further, the constant voltage setting does not need adjustment to account for differences in number or thickness of gels being run.

Find additional tips, troubleshooting help, and resources within ourProtein Gel 1D Electrophoresis Support Center.