contributed by Sean Moore
“Western blotting” is a common name used to describe a technique of detecting specific protein epitopes using antibodies. There are many commercially available technologies for generating a signal after the antibody is bound to the target protein, but our lab primarily uses chemiluminescent reagents and antibodies covalently conjugated to horseradish peroxidase. We use affordable in-house chemiluminescent reagents described in another protocol here.
Gel and Transfer
(1) Resolve your proteins by whatever gel technology you prefer. See Sauer:bis-Tris SDS-PAGE, the very best for my favorite SDS-PAGE technology.
While your gel is running, prepare 4 sheets of 3MM whatman paper with slightly smaller dimensions than the resolving layer of your gel. Also, cut a piece of PVDF membrane that is slightly larger than the paper you just cut.
Soak the membrane in 100% methanol for a few seconds, then pour off the methanol (I reuse mine) and immediately cover the membrane with 1X transfer buffer (below). Don't let the membrane dry out. Place the membrane in buffer on a shaker. Don't wet the papers yet.
Transfer solution 10X “Towbin” buffer:
1.9 M glycine
250 mM Tris
1 mM EDTA (I add EDTA, some people don’t, I find is seems to keep the 10X from turning yellow when stored for long periods)
1X transfer buffer is 1X Towbin and 20% methanol.
(2) Disassemble the gel, cut off the stacking layer, and soak the gel in about 30-50 mLs of transfer solution on a shaker for more than 5 minutes (In my hands, cutting this step causes blotchy Westerns and irregular transfers, free SDS in the gel will compete for protein binding). I soak for 10-15 minutes.
(3) Pour transfer buffer on the papers you cut and bring everything to the semi-dry transfer box.
(4) Assemble the stack. Place two sheets (consecutively) on the bottom electrode. Hold the paper by the edges and "bow" it so the center touches first, then release it. The idea is to prevent bubbles from getting trapped. Carefully place the gel on the papers in the same way. The gel should overhang a bit. Use a razor blade and pinch off the excess gel so its dimensions match the paper's dimensions. Remove the scraps from the box. Now place the membrane on the gel. Try to get it right the first time, proteins on the surface of the gel will immediately stick to the membrane and cause ghost bands in the Western if it's moved. Immediately place two more pieces of paper on the membrane. If you have multiple stacks to make, prepare them sequentially so the membranes don't stay exposed to the air.
Use a smooth cylinder (test tube usually) rinsed in a tray of leftover transfer buffer to squeeze out any bubbles. I place a large crumpled Kim-wipe against the edge of the stack and roll toward it, this wicks away excess buffer than would otherwise squeeze out and short-circuit the transfer when the heavy lid is placed on top.
Using the same Kim-wipe, dip it in some leftover transfer buffer and wipe/wet the upper electrode in the lid where it is going to contact the gel. The jerks in our lab rarely rinse off their buffers from the electrodes so things accumulate on the plates. Also, this ensures a wet contact with the stack.
Place the lid on. You don't need the screws, in fact they will squeeze the buffer out of the gel and cause short circuiting. The manufacturer mentions in the manual that the screws are rarely needed. Connect the power cables and transfer at a constant current of 1-2 mA per cm2 for about an hour.
Blocking and Probing
(5) While the proteins are transferring, prepare a blocking buffer. We used to use 0.2-0.3 grams of BSA or lysozyme in 50 mLs of TBST (expensive). Lately, we almost always use either spray-dried egg white powder, or fish gelatin. Each is inexpensive and we have had the best performance with fish gelatin (it also smells like seagull lunch). Using milk is common and usually fine, but it can impede detection of some epitopes such as His6-tags.
(6) Disassemble the stack by reversing the assembly order. It's easier to peel the membrane off the gel. Before removing the membrane from the gel, pour some blocking buffer onto the blot. This step greatly reduces the smearing of the bands and the "flaming" appearance because it pre-coats adjacent binding surfaces. Transfer the membrane to a plastic tip box and cover with 10-15 mLs of blocking buffer. During these steps, do not let the membrane dry. Even a small bulge from a plastic mold can push the membrane out of the solution and you will get a huge black mess on your blot when you develop it.
(7) After a few minutes, pour off the first blocking solution and cover the membrane with fresh blocking buffer, this step helps to reduce the amount of the free antigen in solution. Shake/rock the membrane in this for 30-45 minutes at room temp. If you're lazy, you can extend the blocking step to overnight at 4 degrees or at room temp in blocking buffer supplemented with 0.02% NaN3 to keep stuff from growing. Some people block overnight at 4 degrees as shown below. Note that this alert researcher let the shaking platform speed up and it rattled his container onto its side, the blocking buffer drained out and the membrane dried overnight, well done. Pro job.
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(8) During blocking, unbound proteins wash off into the blocking buffer. So, pour off the blocking buffer and rinse once briefly with clean TBST before adding your primary antibody. Take note of the corner from which you pour off the reagents, use the same one. If you switch, you can get antigen or antibody re-introduced at a high concentration at the wrong stage.
(9) Prepare your antibody by pouring TBST into the remaining blocking buffer tube and add the antibody (the residual blocking protein is plenty, when using Tween and PVDF, you typically don't need blocking agent during the antibody incubations). The amount of antibody to use can vary greatly, but generally prepare 10-15 mLs at a 1/5000 to 1/20,000 dilution. Shake for 30-45 minutes.
(10) Pour off the primary antibody. Wash the membrane with 2 X 3 5 min TBST changes.
(11) Add TBST containing your secondary antibody. GE recommends using less of the conjugated antibody than usual if you're using the "ECL Plus". I use mine at 1/20,000. Shake for 30-45 minutes.
(12) Wash the secondary off with 3 X 3-5 minute washed, you can go longer on the last few to reduce background.
(13) Develop your blot with an appropriate reagent.