Running SDS PAGE for the first time!

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    • #12697
      dhkwak
      Participant

      Hey everyone. I’m new to this board and joined in hopes of learning various lab techniques.

      I am running SDS PAGE shortly and had a few quick questions:

      Is a running buffer and loading buffer the same thing? If so, are they used as cathode and anode buffers? I have read that cathode and anode buffers are two separate solutions, however some sources suggest that they can be the same solution.

      Also, I am planning on visualizing using Coomassie. Will 50 ug of sample be sufficient for visualization? My protein of interest (BSA) is about 67 kDa in weight and I plan to use a 10T% gel solution.

      Thanks!! Sorry for the very elementary questions. I hope to achieve satisfactory results on the first trial.

    • #97184
      JackBean
      Participant

      Hi, welcome 🙂

      there are several protocols. I have used one with only one running buffer, but last time I have tried other, where were different cathode and anode buffer 😉 Actually cathode buffer can be used in subsequent run as anode buffer (or vice versa?).

      I think, that 50 ug will be plenty of protein. On wiki they write, that 50 ng is enough for Coomassie. Also, you can use several dilutions (depending on how much samples and wells you have;)

    • #97185
      dhkwak
      Participant

      Thanks JackBean,

      I read somewhere that when the running solution contains Tris-Tricine, separate electrode buffers need to be implemented; conversely, when Tris-Glycine is used, the buffer solutions may be the same in each electrode reservoir. I am not sure of the logic for why this is, however.

      I hope this PAGE turns out well. Please let me know if I’m mistaken on anything. Thanks again!

    • #97191
      MrMistery
      Participant

      50 ng should be plenty if you have purified protein, though it also depends – coumassie in my experience can "go bad", especially since most labs reuse that stuff for a million years. But if that’s not the case, you should be fine

    • #97232
      dhkwak
      Participant

      Hey Guys,

      I really appreciate the help. I’m debating as to whether load approximately 50ng or 50ug. I am thinking about measuring concentration of my protein after running it on the gel, however I need to be sure that my protein separates cleanly (although I highly suspect it will not). My analyte is actually a single protein, however it is being cross-linked to a polymer, and cross-linking may occur multiple times per protein. Anyway, I will be using 50ng to start, and if protein separation is adequate, I will be using 50ug to analyze via Bradford assay.

      I have been trying to find more information and protocols (online) for the extraction of protein from the PAGE gel but could not. Can anyone suggest any literature that may be useful here?

      Thanks again!

    • #97236
      JackBean
      Participant

      Definitely do not use 50 ug! That would be much plenty. Just try some dilutions e.g. from 200 to 0.5 ng and you will see, what concentration is fine 😉

    • #97844
      dhkwak
      Participant

      Hey guys,

      I’ve run several gels by now and am starting to feel more and more confident with my technique. However, I am still having difficulty effectively separating proteins of fairly similar MW. In short, I have a protein (BSA) and that same protein cross-linked with a polymer (about 3kDa). The MW of BSA is about 67kDa; cross-linking may occur multiple times per protein but will usually max out at about 10 times (about 30kDa addition, producing a protein w/ a cross-linked agent with MW 67+30 or 97 kDa), generating a range from 67 to 97 kDa.

      I am planning on running a gel soon at 10%. Does anyone have experience with running gels of this range?

      Thanks!

    • #97847
      JackBean
      Participant

      I’m usually running 10 – 12%, but mostly 10%. It’s fine, you just have to take care of the gel not to break it 😉

    • #98162
      dhkwak
      Participant

      I am wondering if there is a rule of thumb for the amount of SDS to use per protein. I have often read that a 4:1 (SDS:protein) ratio is sufficient but am unsure if this refers to concentration, weight, or otherwise.

      Any help would be great, thanks!

    • #98165
      JackBean
      Participant

      SDS should bind in ratio 1.4 g per 1 g of protein.
      Anyway, unless you don’t know the molecular weight of the protein (what if you have mixture?), you can’t calculate the concentration and even have only 4 molecules of SDS per protein, that would be quite insufficient

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