SDS-PAGE is probably one of, if not the most, common techniques in biochemistry. We use it to send proteins traveling through a gel mesh to separate them by size (bigger travels slower) to see what’s inside a sample (based on the # and size relative to a molecular weight standard ladder we run alongside it). 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?! Was it acting as part of a multimer (multiple chains functioning together as a singe unit) or maybe it was simply bound to another protein transiently. With SDS-PAGE you cannot see*!!!! BUT with *native* PAGE you can! (hopefully…) 

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 (there are other methods as well, but this is the one I’m trying so I thought I’d tell you about it. After a couple paragraphs on the blue part the rest will be relevant no matter what kind of native you did). 

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. More practical notes on the blue native at the end

Normal native or blue native, either way, 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. The rest of this is basically relevant to whatever type of native PAGE you started with…

Now you have some options that can basically be split into:

  1. stain (native PAGE as “end of the line” experiment-wise)
  2. transfer to a membrane & use antibodies to probe for proteins of interest (western blot) (native PAGE as “end of separating” experiment-wise)
  3. go onto a second dimension of electrophoresis, commonly SDS-PAGE (native PAGE as the first step of a 2D electrophoresis series) then stain or blot

In terms of option #3… it’s common that after you separate complexes you want to find out what’s in those complexes. To do this you can separate the proteins within the complexes. So basically you cut a lane out of your native PAGE gel (which, remember contains the complexes stuck in place). And you denature those proteins so that the complexes will come apart and the proteins will be able to travel independently from one another, just like in SDS-PAGE. But unlike when you normally run SDS-PAGE, you don’t have liquid samples you can load into wells. Instead, you take that gel slice & And you rotate it 90 degrees and put it on top of an SDS-PAGE gel. (or actually pour the gel kinda around it so it gets stuck in there during the polymerization). 

Instead of the horizontal position being determined by what order you loaded wells in, the horizontal position here corresponds to where the protein (in its complexed form) ran on the native PAGE. So, when you run that SDS-PAGE gel, the horizontal position will correspond to the complex it was in (i.e. proteins in the same “column” will have come from the same complex but there won’t be nice clear separations between lanes because you don’t have lanes!).

The vertical position (how far the protein travels down the gel) will correspond to the size (length here and no longer shape as well) just like you’re used to with regular SDS-PAGE. 

Now you can stain the gel for proteins like normal. But, depending on what your starting sample was, you might see something pretty weird looking with a bunch of spots and smears (you won’t get nice bands because you don’t have lanes! So how clear it looks will depend in large part on how the native run went. 

If you were to draw a line vertically through the gel, the proteins along the line (likely) came from the same complex, so you can then either know what they are based on their size because you started with a pure mix of known proteins or you can probe for them with antibodies specific for proteins you suspect they are. Or you can do things like cut out the spots and send them for mass spectrometry (mass spec) analysis to see their identity (even if you have no idea what they are). 

In terms of option #1 (staining directly after native PAGE)… you can “fix” the gel to precipitate the proteins in place so they won’t diffuse out (wander randomly from where they stopped) before staining or you can stain without fixing and then cut out the bands and get the protein out to identify (with methods such as electrocution or diffusion, none of which I have ever done but there are thorough instructions in the Nature Protocols article I link to at the end). 

*Note: some proteins form multimers through disulfide bridges (cystine cross-links). These are covalent bonds as opposed to just attractions. So they can’t be broken up with just SDS & heat. Instead, you need to add a reducing agent such as BME (beta mercaptoethanol) to split them up. This will reduce the cystines to cysteines, going from |-S-S-| to |-SH HS-|. If you suspect you have multimerization through disulfide bridges, you can run SDS-PAGE under nonreducing (no BME, DTT, etc.) and reducing conditions to see if you see a difference. (and you can do this without having to do the native PAGE step as well). 

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 cuz those proteins are used to being surrounded by lipids. 

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’m added a video 11/16/21 showing me doing it too. Because it’s so cool looking!

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

some good posts/articles:

more on SDS-PAGE: https://bit.ly/sdspagepractical & https://youtu.be/FgXlPAKVSGY 

more on western blotting here: http://bit.ly/westernblotwalkthrough ; video: https://youtu.be/abthP_acYJ0

more on Coomassie staining here: http://bit.ly/cbbgelstaining 


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