Western blots: the what's, why's, and which's, of the workflow

western blots… SDS-PAGE lets us separate proteins by size, but doesn’t tell us what the proteins we separated are. If you add on a second technique, called a western blot, you can use labeled antibody probes to test for the presence and quantity of proteins of interest. It won’t tell you what all the proteins in your sample are, just the ones you test for (kinda like a molecular game of Go Fish!). And you’ll have to have a lot of patience, but it’s a fundamental biochemistry technique, so let’s look at how it works.

The western blot is an experiment used to answer questions like - How much protein X does a cell make under different conditions? The basic premise is - take some mix of proteins (such as the “lysate “you get when you break open (lyse) a cell) → send them traveling vertically though a gel mesh to separate them by size → trap them in place → send them traveling horizontally out of the gel onto a membrane → use antibodies (proteins that recognize specific other proteins) to probe the membrane to see if and how much of a specific protein is there.

Here’s a general layout of the experiment:

SDS-PAGE - separate proteins by size & trap them in a gel

TRANSFER/BLOT - move the trapped proteins to a membrane that likes to bind proteins - kinda like a “protein duct tape.” The membrane doesn’t feel sticky to your fingers but it does to proteins (but not after you’ve touched it with your fingers - so use gloves and tweezers!)

BLOCK - prevent nonspecific antibody binding by getting “generic” proteins to bind the parts of the membrane where your protein isn’t before the antibody has a chance to. Your protein’s only at small portions of the protein duct tape, so you need to coat the rest of that sticky membrane with something “generic” like bovine serum albumin (BSA) or milk (really! it’s chock full of proteins)

WASH, WASH, WASH - wash off non-bound protein

BIND PRIMARY ANTIBODY - Antibodies are little proteins that recognize & bind specifically to specific parts of other things, with the “other thing” being called an ANTIGEN and the “specific parts” being EPITOPES. In a western blot, the antigen is a protein you’re looking for and the epitope is a specific site on that protein. The primary antibody recognizes your specific protein and binds it, so you get your specificity for the thing you’re looking for - yay! but it doesn’t have anything “seeable” about it - boo 😢

WASH, WASH, WASH - wash off non-bound primary antibody

BIND SECONDARY ANTIBODY - this recognizes your primary antibody and has some “detectable” quality like a fluorophore so it will emit light or an enzyme like horse radish peroxidase (HRP) that will convert an uncolored compound you add to a colored compound (chromogenic method) or light (chemiluminescence). It works because antibodies have a unique part that recognizes some antigen and a “generic adapter part” - but that generic adapter part’s only generic for the particular animal that made it (i.e. the adapter part’s slightly different in mice & rats). So you can use things like “goat anti-rat” which is a secondary antibody that uses its unique part to recognize the generic adapter part of a primary antibody made by a rat (and if that rat antibody is using its unique part to bind your protein…)

WASH, WASH, WASH - wash off non-bound secondary antibody

VISUALIZE - detect the detectable using the detectable’s detection method

Some more details: 

continued in comments

[thebumblingbiochemist]

Unlike in the protein purification technique size exclusion chromatography (SEC)(aka gel filtration) where you use pumps and/or gravity to get the proteins to move, & let the proteins run out (and collect them), with SDS-PAGE you turn the power off before the proteins come out so you trap them in place. (you know when to stop it by watching the dye front and/or a pre-stained ladder).

Speaking of the ladder, it’s just a mix of proteins of known sizes to compare to. For “normal” gels we use an unstained ladder (you can’t see the bands until you stain the gel to see all the bands) cuz they’re cheaper but the western blot ones get the fancy one. The proteins in it are pre-stained so you can see them without having to wait to stain them. This will be important for making sure the blot really happened

When you turn off the power, you remove the electric drive helping your protein swim through the mesh, so you trap the proteins in place. The bigger (longer) ones won’t have gotten as far as the smaller (shorter) ones, so they’ll be trapped higher up.



The concentration of protein is high in the gel but low in the surroundings, so the protein can start to diffuse. This diffusion’s slower cuz you don’t have the electric pull helping it overcome friction & gel-tangled-upness but it’s also multi-directional - since their’s no “opposite end” to travel to, it’ll just wander randomly

So you put the gel next to a membrane then change where you put the charge - instead of putting it at the bottom you put it “on the side” to direct the proteins to move horizontally instead of vertically. This is a much shorter trip and you’re not trying to separate any proteins, just move your protein to a safer place to work with it. We’ll talk more on that “Work with it” part later - but first…

It’s not quite as simple as just moving the electrodes (external charge sources). There are different methods for blotting. Older methods include things like capillary transfer, heat-accelerated convectional transfer, vacuum blotting, & diffusion transfer (I thought western blots were slow - but these guys take 2 days just for the blotting part!) Today the dominant method is electroblotting (electrotransfer) which is faster and more efficient.

There are a few types of electroblotting (wet, semi-dry, & dry). I’m going to tell you about WET TRANSFER, which is the kind I’ve always used. Semi-dry & dry are faster, but the transfer’s often not as good

We use a module that fits into the same tank we use for the SDS-PAGE. It’s kinda confusing that in both cases the parts of that get plugged in are both at the top - but that’s just how they get access to the power - the charge they generate is either from the bottom (for SDS-PAGE) or the back (for blotting)

This module fits the sandwich we make, which consists of sponge/filter paper/gel/membrane/filter paper/sponge all wrapped up in a plastic cassette (with holes for buffer to be able to get through) that gets stuck into an “electric oven” - an electrode assembly (blotting module). I called it an oven because it’s somewhere you’re putting the sandwich, but it’s also fitting because it can get hot when it generates the electrical field needed to get the proteins to move. We do the blotting in a cold room. Another option is to use an ice pack

And by wet, I mean it - it’s important to keep everything submerged in transfer buffer when preparing the sandwich. It’s easy for this to get messy but you want to avoid mess-making because the transfer buffer contains methanol, which is hazardous & has to go into the hazardous waste disposal bottle. We usually do the prep in a big glass baking pan

When I first learned how to do a western blot, my teacher was very emphatic about not leaving air bubbles which interfere with the protein’s transfer - proteins can “swim” but they can’t fly! She had us use a glass rod (or a piece of a pipette) to roll out the bubbles. That message stuck with me and let’s just say that, in my first grad school lab experience, I got a little over-zealous with the rolling one time - which happened to be a figure I had to show at a lab meeting during a lab rotation…


Once you get the sandwich set up you turn the power on to generate an electric field to pull the protein out of the gel and onto the membrane (commonly 200mA for 1-2 hrs)

How do you know if the transfer worked? That fancy-dancey pre-stained ladder that we only bring out for special occasions. When you turn off the power and go to remove the membrane, you should see all of the ladder on the membrane and none still in the gel. We can’t see our proteins, but we assume that if the ladder transferred, they did too. Not all proteins transfer as efficiently, however, so you may want to reversibly stain the membrane to check you got good transfer first.

Transfer efficiency depends on a number of things- gel composition, field strength, etc and also protein size -> although they only have to travel through the gel the short direction this time (which is usually just 0.75 or 1.5mm) the same friction gel-tangling-upness applies so it will take bigger (longer) proteins longer to transfer

If all looks good you go from the blot to the block… unlike the antibodies which we choose for their extreme pickiness, when it comes to proteins, the membrane holding the proteins we send the antibodies out probing for is protein-sticky, but it isn’t picky, so you have to take great care to avoid false alarms!

The membrane is kinda like “duct tape” that’s only sticky for proteins - but it’s sticky for ALL proteins. When you transfer the proteins from the gel to the membrane, your protein sticks to the membrane, but only at the spots where it’s present in the gel (the proteins should be moving straight horizontally because the electric field you generate is straight from back to front).

The antibodies you’re going to use are also proteins, so they think the membrane’s sticky too…so if they stuck it’d kinda defeat the whole point of the western - to detect specific proteins. You use a primary antibody that recognizes & binds to your protein, then a secondary antibody that binds to the primary antibody, amplifies the signal, and, because its conjugated (“permanently” attached) to something detectable, allows you to visualize where the primary antibody bound.

[MellifluousLies]

I love your explanations so much! I've only worked in a crazy biotech start-up very very minimal and vague training. Whenever I asked questions, people didn't know or became irritated. 🤐 I really love the underlying science, learning about what's happening in the process, and having multiple ways of viewing and conceptualizing the information! Thank you so much for your videos and website!

Thank you for the detailed explanation. I am an organic chemist and your videos have helped me be able to understand the biochemical techniques that I hear people talking about and to dip my toes into the biochemistry world.

[HandleNigelH]

If i'm using a primary and secondary antibody, can I skip the first wash step after primary antibody binding? I was wondering for a long time why this wouldn't work, afterall, I'll be washing after the secondary ab.

Thank you for all your content btw, i find the depth you go is extremely important for my learning.

[DarkPlayer]

Hello

Why do we quantify the concentration of the protein before putting it on the western blot? I tried searching this up and sources say to standardize how much protein we add to each well but if we standarize it wont it become that all of our bands will be equal and we cannot compare expression between bands?

Thanks