Today, I’m bringing out the big gels! And, although they might look like they’re just scaled-up versions of the SDS-PAGE gels you might be more familiar with, which we use for separating proteins by size, there’s no SDS here – and no proteins – instead, this is a UREA PAGE gel and I’m using it to separate RNA fragments by size.
Both of these techniques are PAGES from the same gel electrophoretic book – the basic premise of gel electrophoresis is that you can make make a gel mesh – in PAGE, this gel is made up of PolyAcrylamide and in the related technique agarose gel electrophoresis (used a lot for looking at DNA) it’s made up of the sugar agarose. They both can make neutral meshes that molecules can travel through, but PolyAcrylamide can give you smaller, more consistent pore sizes, so it’s better suited for looking at little stuff, like the 20-40 nucleotide (RNA/DNA letter) pieces of RNA I want to separate here.
Regardless of your gel-building materials, the principle is the same – the gel mesh acts as a kind of “molecular sieve” – it’s harder for longer molecules to travel through it because they get tangled up more. You can think of it kinda like dropping a jumprope down through a sea of basketball rims. The longer the jumprope, the more it’ll get caught up & the longer it will take to get to the bottom. So if you were to sneak a look partway, it’d be higher up.
In the jumprope dropping analogy, gravity was doing the “motivating” to get the jumprope to persevere through to the bottom – but our gels are too short and the RNA not massive enough for gravity to help us much here – instead we use charge – this is where the E in PAGE comes in – electrophoresis uses electricity to set up an charge gradient through the gel (- on top & + on bottom). Opposites attract, so negatively-charged things will get attracted to the bottom and “fall down,” getting tied up (according to their size) along the way. You don’t let them get all the way through – instead you use tracking dyes to monitor the travel progress and then you stop it when the sizes you’re interested in are in the middle (you can’t actually see the RNA yet cuz it’s invisible but you can compare the dye’s position to what size RNA the dye runs with to get an estimate of where RNA of certain sizes are likely to be)
So – negative charge – with proteins this can be an issue which is one reason to put the S in SDS-PAGE. SDS stands for Sodium DodecylSulfate and it’s an anionic (negatively-charged) detergent (a detergent is a synthetic soap-like molecule that has a long hydrophobic (water-avoiding) tail & a hydrophilic (water-loving) head. One of the reasons you use it when looking at proteins is that it gives proteins a consistent negative charge – depending on their “spelling” (different proteins are made up of different combos of amino acid letters) proteins can have different charges (some of which are +, which would send your protein up not down if SDS weren’t around!
But RNA (and DNA)- regardless of which of the 4 bases (A, U(T in DNA), C, or G) the letters have – has a negative charge thanks to the phosphates in the sugar-phosphate backbone. So they don’t need negative-charge-coating.
BUT the chargedness isn’t the only reason you add SDS when looking at proteins – SDS also serves to denature the proteins – unfold them and keep them unfolded – and do so in a way that keeps them soluble.
You need this shape-eraser because PAGE can only separate things by their length (# of amino acids in proteins or nucleotides in RNA) if they don’t also have different “shapes” – a coiled up jumprope will fall a lot differently than an elongated one. And proteins have shapes – and beautiful ones! I love protein shapes because they can tell us a lot about how proteins work – but for PAGE they make it hard to tell what’s going on in our gel… So we need to unfold them. But we have to do this strategically – the reason proteins fold into the shapes they do is largely so that they can optimize what letters are next to what other letters, hide the hydrophones, let the hydrophiles stick out from the surface so they can play with water, etc.
When you unfold a protein, you expose those water-avoiding hydrophobic residues and they can “panic” and clump together to minimize their exposure to the water. And this can give you insoluble clumps that’d just get stuck in the wells. But SDS – thanks to it’s dual personalities can denature the protein (with the help of heat to loosen things up it can slither in and break up weak (non-covalent) bonds – and it’s hydrophobic tail can glob onto the hydrophobic interior parts of the protein to prevent them from blobbing onto other things.
And since the – charged heads want to hang with water, the protein unfolds, while keeping the hydrophobic parts coated. The protein now also has a uniform negative charge, and because opposite charges repel, the protein gets rod-like.
RNA and DNA don’t have to worry about charge, but they do have shape that needs erasing, thanks to bonds between the NUCLEOBASES of their nucleotides – these bonds can be either w/in the same strand (as is common in RNA) or between the 2 strands of double-stranded DNA or RNA. We don’t usually worry about this for double-stranded DNA – regardless of its sequence, it’s happy in its double helix so even if there’s some shape its the same for all the DNA (see the post about plasmid DNA to see what happens when this isn’t the case) http://bit.ly/2SZbasf
But RNA is often single-stranded and it folds up on itself, forming hairpins and stuff. And, unlike DNA’s double helix, which “doesn’t care” what’s inside, these shapes depend on the sequence of the RNA so different RNAs will have different shapes. So we need to “erase” this, which we can do by adding UREA. Urea replaces the RNA-RNA bonds with RNA-urea bonds, so the RNA strand becomes linear.
So, DNA (linear double-stranded) is good-to-go as is. But RNA needs shape-erasing. But both of these have a built-in consistent charge/length ratio because only their generic backbone part has the negative charge.
So how to do the RNA shape-erasing? There are a few DENATURANTS (shape-removers) we use including FORMAMIDE (in the loading buffer) & UREA (in the gel) – they keep the fragments single-stranded & linear by replacing the letter-letter bonds.
FORMAMIDE & UREA compete for these binding sites, BUT they can be hard to get to (which is part of the point of base pairing in the 1st place – the bases are HYDROPHOBIC (water-avoiding) so they pair (& stack) up in the interior of their 3D shapes & let their HYDROPHILIC (water-loving) sugar-phosphate backbones make contact w/surrounding solvent (more on nucleic acids here: https://bit.ly/2N5sJEW )
To help the denaturants out, we add heat which gives the molecules more energy so they vibrate more & can “pull apart” more easily so the denaturants have easier access. BUT heat’s non-specific – it makes the denaturant-base bonds more likely to come apart too. So we use high concentrations of the denaturants – this way when the bonds break, new bonds will be more likely to form w/ denaturant than w/another base. because this is a “competitive” method, it’s reversible, so we have to include urea in gel itself so there’s plenty of urea the whole run through to keep the bases apart (the hydrophobic (water-avoiding) bases like to huddle together away from water & will happily glue back together if urea’s not there
To provide heat, we heat the sample/loading buffer mixture before loading it & the heat produced by the high voltage provides heat throughout the run. This heat isn’t as high as when we boil it, but the urea effectively lowers the melting temperature (Tm) (temp at which 1/2 the strands have come apart, or melted) so this heat’s sufficient
You also want all the molecules to start from the same place – a nice thing about discontinuous gels like we use in SDS page where you have a stacking & a resolving gel is that the proteins in the well will get smooshed down by an ion sandwich before they enter the resolving gel so if your tip can’t reach the bottom of the well, it’s no big deal. But this is NOT the case with urea PAGE – no stacking gel here so we have to make sure we get the sample to the bottom of the well when we load them. I didn’t discover gel-loading tips until grad school, but they’re amazing 🤩 the long, narrow tips of these tips will help us get to the bottom of the wells (⚠️ but be careful not to go push below the bottom into the gel! ⚠️)
When we run an SDS-PAGE gel to separate PROTEINS by size, we have several options for looking at them afterwards, including colored dyes (often coomassie) & fluorescent stains. With urea-PAGE, we also have several options including 👇
🔸 label & stain-free (UV shadowing)
🔹POST-STAINING – this is very similar to what we you do with commassie & proteins BUT the difficulty is that since the RNA I’m looking at is so much smaller than the proteins I look at, I need a very sensitive stain. A common one we use is the fluorescent stain SYBR gold
🔺pre-labeling: “tag” the RNA BEFORE you run the gel – often by radiolabeling the end w/radioactive (“hot”) phosphate. Afterwards you can expose your gel to a special phosphor screen that can capture a sort of picture of where the radioactivity is. more here: http://bit.ly/35HDDXi
🔸LABEL & STAIN-FREE (UV SHADOWING) – you don’t have to add chemicals, just light! RNA absorbs UV light so if you put the gel on a fluorescent plate & shine UV light – the light will go through the gel, excite the plate, & cause it to give off light. EXCEPT at places where there’s RNA 👉 they prevent the UV light from reaching the plate, so those places show up dark
🔸🔸 especially useful for when you’re running gels for purification – you don’t want to add anything that will contaminate the thing you’re trying to remove contaminants from! BUT you still need to be able to see where the RNA is so you can take it out
But you might have to wait a while before you get a chance to look – these runs can be long (today’s I let go for ~2 1/2 hours) because the gels are often long too – it’s one of a couple of ways we can increase RESOLUTION (what size DNA or RNA fragments we’re able to tell the difference between).
With a longer gel you let the molecules travel further so they have more time to separate – small differences in speed can’t be noticed when you’re not traveling far, but start to make a real “in”visible difference (we can’t see it till we stain it) over long distances. So we can use big gels (often called sequencing gels) to get all the way to single nt resolution. But they’re a pain to pour & take forever to run… Today I was just running these “medium size” ones – but we have tons of different thicknesses and well combs and glass sizes to choose from.
The other way to increase resolution is to change the pore size so that the pores are about the size where the molecules you want to resolve start having a real hard time getting through. If 2 molecules are both a lot smaller than the pores, they can just slide through as if they were the same size, so you can’t tell them apart. But if 1 can slide through fairly easily but it’s a squeeze for the other, the smaller one will charge ahead. But if the molecules are too big for the pores, they’ll get stuck. So we make the gels at a % of acrylamide that’s goldy-loxy for what we want – higher % will give us smaller pores, which is better for resolving small things.
more on casting them: http://bit.ly/2Elhgxa
I have a lot more on the different steps of the process in past posts starting 9/26/18 in the spreadsheet