I usually take my protein sample 0.8ml and sample buffer(2X) 0.2ml for my sample preparation in SDS PAGE. Am I using correct proportion? my protein sample concentration is 4.1mg/ml. What is the standard protocol?

  • 2
    $\begingroup$ Why do you dilute like this? If you do so, the final concentration of your sample buffer is 0.5x. For this dilution you need a 5x sample buffer. $\endgroup$
    – Chris
    Nov 13 '14 at 17:35
  • $\begingroup$ @Chris. Yes, my final concentration become 0.5x. Is it effect on getting bands on SDS PAGE? $\endgroup$
    – kt123
    Nov 14 '14 at 5:24
  • $\begingroup$ Surely it will affect the gel - the concentration of the ions is lower than it should be, so your gel can run funny. $\endgroup$
    – Chris
    Nov 14 '14 at 7:21
  • $\begingroup$ @chris, Can you please give me protocol for preparing sample buffer? Because in different protocols showing different composition. $\endgroup$
    – kt123
    Nov 14 '14 at 9:47

Sample preparation for protein gels is not a complex task. Simply mix the appropriate amount of sample buffer with your sample and load it. For a 2x sample buffer use equal amounts of sample and buffer, for 5x sample buffer use 4 parts of sample and 1 part of buffer (for examle 40µl + 10µl). Heat the mixed samples for 5 minutes at 95°C, cool them immediately on ice and load an appropriate amount of it on the gel. How much is possible really depends on the type of gel you use - 20-30µl should usually be possible.

Do not load too much protein, otherwise you will not be able to seperate them appropriately. I usually use between 20-30µg of total protein per well or as little as 50ng for purified proteins (this depends on the antibodies as well).

Have a look on this webpage for some further information.

For sample buffers I usually use the following recipes. I prefer the 5x, because I can load more protein, if the preparation is not very good. I also prefer the use of DTT to Mercaptoethanol, as it is much less smelly. If you have no DTT (or prefer mercaptoethanol) it can be replaced by an equal amount (200mM in the 2x, 500mM in the 5x buffer). If you use DTT, you need to make aliquots of the buffer and store it at -20°C, a working aliquot can be kept at 4°C for two weeks or so.

2x sample buffer:

  • 4% SDS
  • 20% glycerol
  • 100 mM Tris-Cl, pH 6.8
  • 2 mM EDTA (ethylene diamine tetraacetic acid)
  • 200 mM DTT (dithiothreitol)
  • 0.1 % (w/v) bromphenol blue dye

5x sample buffer

  • 10% SDS
  • 50% glycerol
  • 250 mM Tris-Cl, pH 6.8
  • 5 mM EDTA (ethylene diamine tetraacetic acid)
  • 200 mM DTT (dithiothreitol)
  • 0.25 % (w/v) bromphenol blue dye

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