I showed you how we could use X-ray crystallography to “look” at proteins at the atomic level, where we can see how the different carbon, nitrogen, hydrogen, oxygen, and sometimes sulfur atoms link together into beautiful 3D shapes that allow proteins to act as molecular machines. But usually when we talk about “looking” at proteins in the lab we’re talking about looking at evidence of a clump of unfolded proteins trapped in a gel pore – yup, I’m talking SDS-PAGE! It can’t tell us about a protein’s shape but it can tell us about a protein’s size, purity, and quantity. And it’s a LOT easier than crystallography – so “easy” in fact that it’s a very very frequent staple of life in a protein biochemistry lab. And if you take a biochemistry course, it’s probably one of the first experiments you do. But it can become so “routine” that it’s easy to forget and/or take for granted the awesome science going on at the molecular level.
Today I got to help show the ropes to a rotating grad student. And I’ve become the sort of go-to gel casting tutor. Since I was teaching her, I thought I’d teach you too. So, for today, an SDS-PAGE review!
Sodium DodecylSulfate PolyAcrylamide Gel Electrophoresis is one of the most frequently used techniques in biochemistry. We use it to separate proteins by their lengths by using electricity to send them swimming through a gel mesh.
Longer proteins get tangled up more and travel more slowly, so when you stop their swimming they won’t have traveled as far and when you stain the gel to show their location they’ll be higher up the vertical gel slab. For this protein character counter to work, your “input file” (protein) has to be in the proper format (unwound and negative-charged), so let’s take a look at how we do it!
Proteins have different lengths, shapes and charges, so if you want to separate based on length alone you need to erase the shape (denature them) and “paint over” the natural charges with an even negative charge (you don’t want to just erase the charge cuz you need a negative charge to motivate them to move through the gel towards the positive end).
Just like all poems are written using letters from the same alphabet but they can be dramatically different, proteins are all made up of letters (amino acids) from the same protein alphabet but, depending on what combinations of letters are there, they can have wildly different properties and are thus best suited for different purposes. Would you read the Iliad or the Odyssey to a group of preschoolers? Or Cat in the Hat to college students (I hope so on this one :P).
Also like poetry,
What if you want to know how long a poem is (how many letters it has)? As we saw above, you can’t just “count the lines” (shape & size can have different relationships). Instead you need to get all the poems in a single long line format (e.g. proteinscantakedifferentshapes). This requires unfolding the protein, removing its “natural shape” and hence “denaturing it”
A key thing to note is that all of these letters (even if they look like they’re on “different lines”) are strung together LikeCursiveLetters through strong peptide bonds in their generic backbones. But different letters also have unique “side chains” (aka “R groups”) sticking off with different properties (small, big, +, -, etc.). We looked at each of these through #20DaysOfAminoAcids http://bit.ly/allaminoacids
And speaking of charge… we’re going to use the “opposites attract” strategy to motivate proteins to move through the gel. But proteins can have different charges –> certain letters can be + or – charged and different proteins have different numbers of those letters (e.g. imagine that the dots on i’s are + charges & the – on t’s are – charges. since different poems have different numbers of them, they’ll have different overall charges). And this can “confuse” our character count apparatus.
LENGTH is what determines how fast they move (& thus how far they’ll have traveled when you stop the experiment) but CHARGE is the motivating force that makes them even try to move in the 1st place. So you need to make sure that charge & size are *directly* related
So we need to remove the shape, keep it removed, and “standardize” the charge so that all the proteins are – charged (and thus motivated to move towards the + bottom of the gel) and that charge is directly related to the length of the protein not to the identity of the letters. So what kind of “erasers” and “charge paint” do we need?
Let’s start with the shape erasers – The shape comes from different letters interacting with each other (e.g. those negative “crossed t’s” wanting to hang near the positive “dotted i’s”) and with the liquid (often water) they’re bathed in (e.g. water-loving (hydrophilic) letters wanting to be on the surface & water-avoiding (hydrophobic) residues wanting to stay hidden in the core). But the more energy the letters have, the more likely they are to explore! And heat is a form of energy, so we can raise the temperature to denature the proteins.
Imagine protein letters interacting as 2 people shaking hands. At low temperatures, it’s more like they’re holding hands. But adding heat is like getting them to shake really vigorously. This makes it harder to stay held, so they come apart. But if you don’t do anything, and you lower the temp again, the hands can reclasp, refolding the protein – or even worse they can reclasp with a new partner, changing the protein’s shape – often in nonideal ways that makes the protein clump up (aggregate).
So, once the hands separate, you want to coat them with a slippery soap. True story – one time when I was a little kid, I thought I got locked in a bathroom when really my hands were just not rinsed well after washing them so the soap made the door handle too slippery – SDS (sodium dodecyl sulfate) is a detergent (artificial soap) and it works kinda like that. Once the protein letters declasp, SDS slithers in to coat them and prevent them from reclasping
So heat and SDS both work in the shape-erasing part and SDS plays double-duty because it also does the charge painting! The reason SDS is able to slither in and coat the protein is because it has a long hydrophobic tail (the dodecyl part) that globs onto the hydrophobic parts of the protein that are normally hidden in the core. If you unfolded a protein but didn’t cover those parts, those parts would panic & clump up in aggregates to minimize water exposure, thus making them insoluble -> but the SDS isn’t all hydrophobic – as an “amphiphilic” molecule it has hydrophobic AND hydrophilic parts – it has a – charged (anionic) head group (the sulfate part) that likes to hang out with water, so it keeps the coated proteins soluble. And it doesn’t like to hang out with other – charged heads so it keeps the protein unwound and “rod-like” – ideal for slithering through a gel.
Those – heads don’t like each other but they DO like the + charge that forms at the bottom of the gel when you turn on the power to create an electric field. The electric field is created and maintained through electrolysis, which I won’t bore you with here, but you can learn more about here: http://bit.ly/2XMLMbG
Now that we’ve seen how we prepare the protein let’s look at the gel we’re going to send it through. The gel is made up of polyacrylamide and cast into thin vertical slabs with wells at the top where you put in the protein. The motivating + will be at the bottom, so the proteins have to go through the gel if they want their reward.
But the journey isn’t easy – especially if you’re big! Polyacrylamide is neutral & non-reactive, so it doesn’t *chemically* sidetrack the proteins on their journey, but the mesh runs *physical* interference, acting as a molecular sieve. Bigger molecules have a harder time squeezing their way through mesh’s holes. They experience more frictional “pushback” from the gel & travel more slowly. If you have a really loose mesh, small things can slip through easily (& thus travel together, “unseparated”) & only bigger things get held up, but if you tighten the mesh, smaller things start getting slowed down too.
We take advantage of this to get all the proteins to the starting line at the same time. Depending on your sample volume & well size, the proteins might be very spread out vertically in the wells when you start the run. If we didn’t do anything about this, the protein molecules that happen to be closer to the well bottom would have a “head start” so if the molecules traveled at the same rate, molecules of this same protein that were higher in the well after loading your samples would lag behind so you’d see multiple bands for the 1 protein (probably more like a smear). Also, molecules of different sizes traveling at different rates but from different starting points could end up at the same position at the end of the run.
To make the race fair, we use “discontinuous” gels – they have 2 layers of gel stacked vertically. The main, bottom gel is the resolving (running) gel, & on top of that of the resolving gel is a “shorter” layer of “stacking gel” (this is the layer that will have the wells).
The stacking gel has bigger pores so the molecules travel quickly until they reach the barrier w/the denser resolving gel that slows them down so you get a “bottleneck” where the proteins pile up into a single concentrated band. So all the protein molecules start at the same “starting line” & enter the resolving gel at the same. It might help to think of this as like a track race. A continuous gel would give you a sort of “mass start” whereas a discontinuous gel allows all the molecules to start at the starting line.
How do we make those pores bigger? By using less polyacrylamide and/or less crosslinker -> polyacrylamide is made up of chains of acrylamide. And we make it meshy by bridging those chains with a cross linker, bis-acrylamide. So if we alter the amounts we add, we can manipulate the pore size, so mesh tightness is often reported by % acrylamide and/or crosslinker. day You can also adjust mesh tightness of the running gel to get a looser mesh if you want to separate bigger things or a tighter mesh if you want to separate smaller things.
A lot of labs buy pre-made gels and if you don’t use them a lot, that can be worth it but if you use a ton like we do it makes a lot more sense $$-wise to make your own (and it makes you feel less guilty about running 15-well gels when you only have a couple of samples!). Plus, when you make your own you can customize to your tastes (not just meshiness but also thickness, # of wells, etc.). To make them, you mix the acrylamide & bis-acrylamide, add polymerizing agents and pipet it in between glass plates before it gelifies. more on this here: http://bit.ly/2NkBoUt
so, prepared protein: check
prepared gel: check
Now we just need to put the E in the pagE! We introduce the SDS-coated protein to the gel by sticking it in wells at the top. But to get the protein to actually move through the gel we have to turn on the (properly connected) power. This is where the ELECTROPHORESIS part of gel electrophoresis comes in/ We immerse the gel in a salt water bath with a ➖ electrode (cathode) at one end & a ➕ electrode (anode) at the other end. This separation of charges generates an electric field
If you put something charged in an electric field, it’ll move towards the oppositely charged end. We run NEGATIVELY-CHARGED molecules towards the POSITIVELY-CHARGED anode (run to red) ➖➡️➕ And, thanks to the SDS, the proteins all have a negative charge that’s directly proportional to their length, which serves to “standardize” the charge so separation can occur by length.
So the proteins start slithering through the gel and, kinda like dropping a jump rope through a sea of basketball hoop rims, they’ll get tangled up, with longer proteins getting tangled more and traveling more slowly. When you turn off the power (which you know when to do by watching a tracking dye) they’ll lose their motivation to move and stay stuck.
You can see that tracking dye, but the proteins are invisible to the naked eye! So to see them you need to stain them. A common stain is commassie brilliant blue (CBB) which is a “total protein stain” that’ll turn your proteins (wherever they are) blue, so they appear as bands in the gel. more here: http://bit.ly/2ElRed7
Each protein will appear as a band – the vertical location tells you about the protein’s size (which you often compare to a molecular weight “ladder” of proteins of known size). The thickness of the band tells you about how much protein there is. And the number of bands tells you about the purity of the sample (more bands, more proteins (but beware of multiple proteins of similar size running together and thus appearing as a single band!).
During a protein purification, you start with a mixture of the protein you want and whatever else the cell making your protein made for itself – and you want to see extra bands disappearing until you see a single strong band at the end!
If your protein biochemist friend says they can’t go to the movies because they have to “run a gel,” SDS-PAGE is probably what they’re referring to. If your molecular biologist friend gives you the same excuse, they’re probably referring to an agarose gel which is used to separate DNA – I use those too (in the molecular cloning phase when I’m trying to get a gene for my protein into a plasmid to express it from) and you can learn more about it here: http://bit.ly/2SDKE8I
but in my opinion, SDS-PAGE is the cooler molecular ruler!