From time to time, I will give a glimpse into the “glamorous” life of a research associate and talk about what I’m doing in the lab on a particular day. These entries I will call “A Day in the Life…”
Today I’m doing a Western. No, I’m not running around the lab in a cowboy hat with pipets in my holsters. A “Western” is what we lab rats call Western blot analysis, a technique for separating and detecting proteins.
The first step of a Western is to get some proteins to analyze. In my case, I will be using proteins found in cultured cells. The cells have to be opened up so the proteins within them are available for the assay. This involves “popping” the cells open using a chemical detergent to break open the cell membranes. The end result looks like a somewhat cloudy liquid.
The tube that I'm holding contains "cell goop" or as we call it in the lab "cell lysates." This sample also has loading buffer in it which gives it the blue hue.
Next, the cell goop is placed into a buffer that will help “sink” the proteins into a thin rectangular “well” that is part of the separating gel. The gel is extremely thin and is smooshed between two glass panes to help hold it in place. The gel sort of has the consistency of Jello – a really, really thin sliver of Jello.
This is the gel which is wedged between two plates of glass. The green comb was placed in the top of the gel to form the "wells" into which the sample will be loaded (once the comb is removed, of course).
The comb is removed and the gel (still smooshed between the glass plates) is placed into the gel holder. The two metal posts on the top will allow electricity to flow through the gel, once the apparatus is completely assembled.
Here I am loading the sample into a well on the gel. The sample is inside the pipet tip which resembles a hollow tube; the tip is attached to a pipeter which very accurately measures how much liquid sample to draw into the pipet tip.
Once the cell goop (or protein samples, as we call them) are loaded onto the gel, we apply an electrical current conducted through liquid that has a few goodies in it (goodies like detergent, the amino acid glycine, and more). The proteins separate based on their size – the smaller proteins wiggle through the spaces in the gel and travel farther than the larger proteins. This is kind of like a bag of potato chips – the larger chips stay near the top of the bag, the broken chips settle somewhere in the middle, and the crumbs settle at the bottom.
Here the gel and loaded samples are in the Western blotting apparatus on the right. A 2-milliAmperage charge is applied to the gel using the power supply on the left
This part takes two to three hours (at least in my lab it does). So there’s some time to do lab chores or other experiments while the gel is “running.” A watched gel does not run faster.
This is how the protein standard looks at the start of the "running" process. Note that all you can see is a band of blue indicated with the yellow line I added.
This photo was taken an hour or so into the "run." Note that instead of one band of blue seen in the previous photo, you now have several bands (here seen in two "lanes" -- as the contents of the well runs from the top to the bottom of the gel, we call those "lanes"). You should be able to see some blue bands and at least one red band. The protein bands closer to the top are larger in size than the protein bands that are closer to the bottom.
How do we know when it’s done? Well, in one of those wells, we loaded a “protein standard” which we buy from a commercial source and contains specific proteins of specific sizes and are colored with specific colors so that we can see the proteins as they separate. When the smaller proteins in this standard get close to running off the gel (at the bottom), that’s when I stop my gels. Other researchers may want to separate the larger proteins more, so they might let some of the standard proteins run off the gel before stopping the separation.
Once the proteins are separated on the gel, we transfer those proteins to a piece of specialized paper, in my case it is called nitrocellulose. An electrical current moves the protein out of the gel and onto the nitrocellulose. This takes about an hour, so not as many lab chores can be done while waiting for this step.
Here I am transferring the extremely thin gel by hand to another apparatus. The gel is very thin and flexible as seen here.
Here the gel has been placed on some paper. You can see the protein standards (the colored bands on either side). The actual protein samples are between these two colored standards -- you can't see them yet because they are colorless, but they are there.
This is what the transfer apparatus looks like before I put the lid on it. Again, the two metal posts will conduct the electrical current through the gel toward the nitrocellulose membrane. The proteins which are in the gel will be "transferred" by this current to the neighboring nitrocellulose membrane.
The transfer set up now looks very similar to the running set up -- the difference is the part that's inside the green-topped box on the right. The power supply on the left delivers 47 volts for one hour to complete the protein transfer.
This photo was taken after the transfer process. You can see that there are no proteins left in the gel (bottom). The protein standards can now be seen on the nitrocellulose membrane (top).
This nitrocellulose containing the transferred proteins is called a “Western blot.”
Here are the proteins on the nitrocellulose membrane. The membrane behaves a little like a piece of paper.
The outside edges of the nitrocellulose membrane are trimmed, a corner is notched (upper right) so that I can tell which side is up.
From here it goes into a little plastic container. Non-fat dry milk is added and the membrane is incubated overnight in the lab fridge. End of Day 1.
But wait there’s more! Just having the proteins on the nitrocellulose isn’t our stopping point. We want to see if a certain protein we’re interested in is present in the samples on the blot. But first we put the blot in some blocking solution – usually it is a solution of non-fat dry milk (yep, the same stuff you find on the grocery shelves).
We have to block the spaces on the blot between the separated proteins on the blot so that we detect only the protein(s) of interest and not the empty spaces on the blotting paper. The next step is to add another protein, called an antibody, to the blocking solution. If the empty spaces were not blocked with milk proteins, then the antibody would non-specifically stick to those areas on the blot – we call this background signal and we want as little of that as possible.
The antibody is designed to stick only to a certain protein, which hopefully is present on our blot. We allow the antibody to mix and mingle with the blot, using some gently shaking, giving the antibody a chance to find and stick to the protein if it is there.
After washing the blot to remove any antibody that hasn’t stuck, we add another antibody that will seek out the first antibody. This second antibody has an enzyme stuck to it so that when we add the right chemicals, the enzyme will react with the chemical and produce a tiny bit of light which we can capture on a piece of film.
Day 2. Most of the day is spend rinsing the blots. First round of rinses involve dumping out the blocking + specific antibody solution and adding a buffer containing salt and detergent which helps remove any antibody that has not stuck. This step involves 4 cycles of rinsing, each cycle is at least 15 minutes.
Each 15-minute rinse step involves agitation. Here the blots (in the plastic containers) are gently swirled using a rotary platform. They look a little blurry because of the camera exposure setting.
More rinsing! This is affter the secondary antibody has incubated for half an hour with the membrane. Again, 4 cycles of 15-minute rinses.
Next step is in the dark room, and yes, we use a red light in the dark room. This is the film processor. The blots are placed in a special buffer that reacts with the tag on the secondary antibody. This reaction generates a bluish glow at the exact spots where the antibody is stuck -- often you can't see this with your eyes, but sometimes you can. The blot is briefly exposed to a piece of special X-ray film and the film is developed in the film processor (shown above).
If all goes right (and quite a bit can go wrong along the way – we call that troubleshooting when we have to figure out what went wrong), we will see the protein as a little black rectangle on the film.
Here is the result. On the left is the original nitrocellulose membranes (wrapped in plastic wrap so they don't get the film wet). The film is on the left. The black "bands" are where the light from the reaction with the antibody that was stuck to the protein of interest hit the X-ray film. Now all I have to do is match up the film with the blots (so I know how they relate to the protein standard sizes) and I'm done. The blot and film are sitting on a light box so that the film is easier to see (like an old-fashioned X-ray film).
Now you know one way scientists can detect proteins. (Note: there are many ways to detect proteins and I’m sure I’ll be telling you more of them in the future.)