Hi,

At high concentration (1-2%) the published saturating SDS:protein binding ratio is about 1.4:1 by weight, that is roughly one SDS molecule per two aa on average. It is dense but  not that dense to prevent any further interaction.  More importantly, as a quite hydrophilic small molecule SDS should have no trouble dissociating from the peptide when its in-solution concentration drops (therefore you can use SDS gel bands for MS). With common procedure, during staining the SDS should partially fall off( especially if the gel is heated), and partially remain with the protein in the gel, depending on: how hydrophobic the protein is, how low the environmental SDS concentration becomes, how much organic solvent there is in the solution, etc.. The coomassie should simply find whatever hydrophobic/positively charged patch to bind and aggregate. Besides, coomassie-R is probably slightly more hydrophobic than SDS so it is capable of competing SDS off if necessary. (The even less hydrophobic Coomassie G250 definitely binds protein in the presence of detergents - that how Blue Native gel works for membrane proteins) Finally, since the only thing you are looking for is some deeper blue to indicate the presence of protein, even if SDS did prevent dye binding to some extent, your gel still will work. This is different from when you want to use the dye to do some quantitative work such as the Bradford assay. It would be interesting to know the effect of detergents on Bradford. 

Zhijie

On Oct 4, 2018, at 9:26 PM, Alex Lee <[log in to unmask]> wrote:

Dear All,

I am thinking that in an SDS-PAGE experiment, if protein samples are boiled in SDS containing loading dye, and supposedly SDS interacts with proteins, why the Coomassie Blue dyes could still interact with and stain the proteins?      I am thinking SDS is covering the proteins, making no room for the Coomassie Blue dyes interaction.  I'd appreciate it if any input from this forum.

Alex

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