Blue Native PAGE (BN-PAGE) behind the scenes

Native PAGE lets you separate proteins in their native (unfolded) state in a PAGE gel, keeping complexes together unlike SDS-PAGE. And the “Blue” part uses a negatively-charged (and pretty!) dye to allow even non-negative, “basic” proteins to run!

For more on the theory, see https://bit.ly/nativepageoverview ; YouTube:    • Native PAGE (PolyAcrylamide Gel Elect...    

When we add SDS, we lose a lot of information about proteins. And this is part of the point. SDS is a detergent that, along with heat, helps denature (unfold) the proteins and coat them in a slippery negative coat to keep them soluble. With SDS-PAGE we want to remove a protein’s “shape” in order to separate it just by its “length” (i.e. how many amino acids are in its chain). And SDS is great for this. But, what if we wanted to know whether that protein chain was alone?! With SDS-PAGE you cannot see*!!!! BUT with native PAGE you can! (hopefully…)  
 
blog: https://bit.ly/nativepageoverview ; YouTube:    • Native PAGE (PolyAcrylamide Gel Elect...    
 
In native PAGE you do NOT denature the proteins. You just run them as is. This way multimers will stay multimers & complexes will stay complexes (if they’re strong enough) so they’ll run like a bigger thing (sum of their sizes, except here you also have shape to contend with so the size vs run speed relationship is less clear. But the importance of SDS before wasn’t just to denature and prevent aggregation, it was also to give the proteins a negative charge. They need that negative charge in order to be attracted through the gel towards the positively charged electrode. And not all proteins are negatively charged at the pH you run gels at. Some are, so you can just run them same as usual, just don’t add SDS or dye, just add a little glycerol to your samples to keep them from floating out and run them in the cold room with less intense voltage to keep the proteins stable. BUT if your proteins are not negatively charged, if they’re too “basic” they won’t be tempted to travel through the gel and instead will get stuck in (or even travel up out of) the well! This is where a technique called blue native PAGE (BN-PAGE) can come in.
 
The “blue” in Blue Native comes from the Coomassie Brilliant Blue dye that is used (yup, just like the dye you might use to stain your gels for protein or determine concentration with a Bradford assay. The dye (you use the G-250 form) is anionic (negatively-charged) and can weakly bind to proteins, thus making the proteins negatively charged and giving them the charge needed to go. In this way, it’s similar to SDS. BUT the dye is NOT a detergent and it’s not denaturing. So it will kinda just gently stick to your proteins, hopefully without even disrupting any complexes. Because this reaction is so weak, however, you need to put dye in the running buffer so that if a dye molecule comes off during the proteins’ journey there’s plenty more to take its place.  
 
End result is that your proteins end up traveling through the gel no matter how basic they were.
 
The proteins’ travel speed will depend on their size & shape (and remember here we’re talking about at the complex level). Typically you use a gradient gel so the gel’s mesh is looser at the top (good for separating big things) and tighter at the bottom (good for separating little things). The proteins and complexes can then travel through the gel until it gets too tight for them to move much further, which will depend on their size.  
 
When you turn off the power source, removing the charge gradient, the proteins will stop moving.

note: BN-PAGE is commonly used for membrane proteins, so if you look up protocols they often talk about solubilizing membranes and stuff and include a mild detergent  
 
Blue Native PAGE (BN-PAGE) - here are the notes I took for my lab book, but there are better protocols linked below. There are lots of different protocols and you can use different gel types and stuff but this is how I (just) learned.
  
I used the Biorad 4-20% Tris-Glycine TGX gels 
 
Prepare your samples and instead of SDS loading buffer, use glycerol to final concentration of 20% (I started with an 80% glycerol stock because it’s much easier to pipet than pure glycerol! So, 80%’s my 4X loading buffer in a way… and the glycerol keeps the samples from running away.  
 
DO NOT BOIL! Boil in SDS-PAGE to denature, but here you do not want to denature 
  
Set up your cassette in the running module 
 
pour no-dye buffer (1X Tris-Glycine) into inner chamber (cathode buffer) 
 
load wells 
  
load 20% glycerol in empty lanes to get consistent flow 
  
add dye to inner chamber - pipet 2mL of 100X concentrate into the bottom of the chamber, then pipet up and down with a large pipet to mix 
  
run for ~20 min at 100V - proteins should enter gel 
  
remove inner dye-containing buffer & replace with fresh no-dye buffer 
  
continue running until dye front comes out - can also increase V to 200 
 
detailed protocol:  https://doi.org/10.1038/nprot.2006.62