Native PAGE (PolyAcrylamide Gel Electrophoresis) & Blue Native PAGE

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…)
blog form with full text & figures: bit.ly/nativep...
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

[@DoctorJammer]

I love that you show the actual lab setting. Gives me a better frame of reference.

[@thebumblingbiochemist]

Glad you find it helpful!

[@lohanemoreira2590]

This is the best explanatory vide I've ever seen!!!!!!!!!! THANK YOU SO MUCH!!!!

[@thebumblingbiochemist]

So glad it was helpful!

[@anisheshdas362]

Thank you. Reallh helpful.

[@thebumblingbiochemist]

Glad it was helpful!

[@mengda9161]

Great post. This is just what I need.

[@thebumblingbiochemist]

Happy I could help!

[@yazallen8878]

Awesome channel! Thank you so much.

[@thebumblingbiochemist]

Thanks so much!

[@roidedv480]

thanks for sharing！this helps me a lot

[@thebumblingbiochemist]

Glad it helped!

[@kaydenkwah5280]

OMG you're AWESOME!!!! What a lifesaver :D

[@thebumblingbiochemist]

Thank you! So glad it was helpful

[@spectator59]

Have you tried native PAGE with proteins statically bound to nucleic acids?

[@thebumblingbiochemist]

as in an EMSA? I have content on that

[@spectator59]

@@thebumblingbiochemist Yeah, EMSA's close.