Nope, not those kinds of westerns.
THESE kinds of westerns are for looking at proteins! Technically they’re called western blots. There’s also such things as northern blots, southern blots, and eastern blots, but I’ve never done them myself.
Western blots like these ones let me see specific proteins of interest in any given sample. For instance, I could have whole bacterial cells, but I might have a very specific, single protein I want to make sure is contained in those cells.
To check that the protein is in my bacterial cell, I can break open the cell by putting my sample in a buffer with some very strong detergents and chemicals, boil them, and then separate them out on a protein gel using an electrical current. This part is called SDS-PAGE (SDS is the detergent, PAGE stands for polyacrylamide gel electrophoresis).
The protein gel is like a matrix or a mesh. When you load your protein samples at the top of the gel and run an electrical current, the detergent mix makes the proteins slightly negatively charged, meaning they travel to the positive terminal at the bottom of the gel at various different speeds, depending on how big your protein fragment is.
For instance, if your protein fragment is huge, it has a harder time getting through the gel matrix, meaning it’ll travel slower. If your protein fragment is teeny, however, it will travel faster through the gel and end up closer to the bottom of the gel.
If you stop the gel before all your protein fragments run off it, you’ll get a nice separation of your proteins within the gel.
The next part is the actual western blotting component. You need to actually transfer your proteins from your gel, onto a special membrane that can absorb and retain proteins.
How would one do that? By applying yet another electrical current.
You basically make a sammich of your gel, sitting on top of your membrane, in between two electrodes. When you run an electrical current, the slightly negatively charged proteins move toward the positive terminal, and get absorbed by the membrane. By the end you’ll get all your proteins sitting on your membrane like you can see below, where the protein ladder is visible.
Each of those coloured bands are proteins of a specific, known size, and you can use that to compare against your protein of interest, in order to determine whether the protein you can see at the end of this whole process is of the correct size and is actually the one you’re after.
You can’t see my protein there, yet. I need to perform a series of steps before I can actually visualise it. It’s there, but in its current form it’s invisible to us.
So, the first thing I’ve done with my membrane is to ‘block’ it. Usually in a mix containing buffer, detergent, and a non-specific protein of some description. In my case I’ve used 5% skim milk, because having full cream milk can interfere with the protein binding (apparently the fats are bad). You can also use bovine serum, too. Just depends what’s more convenient. Our current bovine serum powder ends up looking like pee, which makes me giggle a bit.
The blocking step ensures that any part of the membrane that currently hasn’t got any protein on it, absorbs (in this case) the milk protein. The membrane is very absorbent for proteins, which means any part of the membrane that remains unblocked and exposed could jeopardise the next step, which involves adding antibodies.
For those that aren’t aware, antibodies are also proteins. They’re typically quite good at targeting a specific ‘antigen’ (target protein fragment), so we use these to select out a protein of interest. Say I’m looking for Protein A. After the blocking step, I’ll add antibodies which bind only to Protein A.
But- if I didn’t block my membrane beforehand, my antibodies will just get absorbed onto the membrane itself. It’ll stick around, not because it’s found Protein A, but because it’s stuck to the membrane. This isn’t good for me and my experiment. Any interactions the antibody has, needs to be to its target protein, not the membrane. Otherwise I’ll get false results or a really messy blot that looks like dark storm clouds.
You need to wash after adding the primary/first antibody, to make sure any antibodies which haven’t bound its target protein is removed. For some reason the magic number is three washes. They could be three 5 minute washes, 10 minute washes, three ‘oops I left it for an hour because I forgot my experiment’ washes… Either way, a good rinse in buffered saline and detergent is required.
You then come in with your secondary antibody. This antibody now binds to your primary antibody (in my hypothetical scenario this would be the antibody I used to bind Protein A). You essentially want to create another sandwich stack, which ends up looking like the left panel…
You can get a scenario on the right, where the primary antibody is bound (conjugated) to an enzyme (denoted as ‘E’ in the diagram) or such like, but that’s a very expensive set up so typically we have the situation on the left.
Either way this second antibody is bound to something, it could be an enzyme which, when given its substrate (the compound that it works on), it produces a colour, a chemical which can be detected using a special machine… or, it could shine or fluoresce when you bombard it with lasers… either way, it now makes your protein visible.
This is a blot I made earlier in its finished form. I’ve got my molecular weight/size marked out on the left (the unit we use is kilodaltons or kDa), based on my coloured protein ladder. I’ve got my five bacterial strains specified at the top. The primary antibodies used are specified (ie. what the hell are the proteins I’m showing?), and then the bands for the specific proteins of interest. I had to use a special machine to actually visualise the proteins, because otherwise they’d just look like the blot from earlier that was sitting in milk.
Or you could get a scenario like this…
Which is… accurate. That’s actually been the last 48 hours. Currently trying to make it better.
Ahh, Sketching Science, you’ve done it again.