I’ve written before about western blots (I’ve written an overview here), but as I was running what may become my last protein gel, I thought I’d just document it and write a lengthier post on them.
So as I’ve written before, we use western blots to visualise proteins. I’m currently checking some Coxiella strains to make sure they were making the proteins I wanted them to.
In order to look at individual proteins, the first thing that happens is that I collect my samples and place them in a really harsh detergent. The components of this detergent mix kill and break open cells, then turn proteins, which initially look like squiggly blobs…
… into nice protein fragments. Because proteins have specific conformations and shapes, but for the kind of protein gel I’m running, I need them to all be linear. This is called ‘denaturation’. The detergent mix also binds to the proteins and make them slightly negatively charged.
So I add my detergent mix, boil my samples for 10 minutes at 95 degrees, and then load them onto protein gels, which in fancy terms is called sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE). Polyacrylamide in liquid form is highly, highly toxic, so you have to be careful and wear proper protective equipment. Fortunately for us, we have pre-cast gels that are already solidified and ready to use.
Once I’ve placed the gels in the gel tank with the relevant buffer (it has some electrolytes in there so that an electrical current can pass through), I can begin loading my samples. You can see how teeny those individual wells are. These wells in particular hold 20 microlitres of sample (a microlitre is 1/1000 of a millilitre- i.e. one thousand microlitres makes one millilitre).
Once the samples are loaded, alongside a molecular weight ladder that has known proteins (with known sizes) in it, I can run a current, and the gel will run.
As mentioned, the proteins are bound by the special detergent mix, and are slightly negatively charged. When the current runs, they slowly move toward the bottom of the gel tank, which contains the positive terminal.
Large proteins have a harder time moving down the SDS-PAGE gel matrix (kinda like mesh), so they move slower. Smaller proteins can move through the matrix quicker, so they move faster down the gel. Once the samples have had a chance to move down the surface of the gel, all the protein fragments will have separated out according to size. You can tell that a sample is running by looking at the detergent mix in the samples, which in this case is blue. The red is from something else in this experiment, and is actually just a funky addition.
Once the blue colour has reached the bottom of the gel, you can stop the current and take the gel out. At this point the protein gel/SDS-PAGE needs to be transferred onto a special membrane, which turns it into a western blot.
To do that, I use a spatula to break open the gel cassette.
You can see how thin that gel is.
I have a transfer stack ready to go so that when I take the gel out, I can put it straight onto my special membrane. The special membrane looks like weirdly chalky paper, but it’s quite cool because this stuff is highly absorbent for protein. If you put protein on it, it will soak it up like a sponge.
So once I take my gel out, I can plonk it on to my membrane. Currently the proteins are still in the gel itself.
In order to move the proteins from the gel to the membrane, I once again need to run an electrical current through my gel and membrane.
So at the bottom of this ‘sandwich stack’ of membrane and protein gel, there’s a layer of filter paper and an electrode pad. You’ll see an example set up illustrated in my previous post.
Then I add a layer of wet filter paper on top of my gel, and gently squish the whole thing with a roller to make sure I don’t have any air bubbles. If I have an air bubble between any of these layers, then the electrical current won’t pass through that area, and my protein sample won’t transfer from my gel on to my special membrane.
Once I roll out the filter paper, I then add an electrode layer to complete my sandwich stack, and I’ll repeat the process of removing air bubbles. The whole stack (aside from the filter paper I add on top of the protein gel) is already semi-wet (soaked in electrolytes).
There’s an extra layer in this kit to connect the top layer of electrode to the machine itself (that’s what that white sponge is on top).
Once it’s sitting in the machine, I can close the lid and run a current. The whole process for us (because we use a pre-made kit), takes 7 minutes. The old school way (no kit) takes about an hour.
Once the transfer is complete, it’s the moment of truth…
You can see now that the molecular ladder is no longer visible in the gel, but is sitting in the membrane. That’s good! That means your proteins (currently invisible) are also embedded on the surface of your membrane.
I’ll now stick these delicate membranes in a mixture of buffered saline (keeps the proteins happy), detergent and skim milk powder. If you have a look a the previous post, you can see that this step is called ‘blocking’.
I’ll add the primary antibody after 1 hour of having these membranes gently rocking on a rocker, and leave it overnight in the cold room (also gently shaking). Hopefully my proteins were being made by the bacteria, and my primary antibody can find and bind them. Thus begins the steps to actually visualise my proteins of interest!
Categories: Ph D posts
A former wet-lab based Bacteriology Ph. D student residing in Australia. Now working part time at a secret location as a Communications and Data Officer. 👀 🦠 🧫 🧬