Sauer:Western blot

Western Blotting
contributed by Sean Moore

Background
“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 uses primarily the “ECL Plus” chemiluminescent reagents from G.E. Healthcare and antibodies covalently conjugated to horseradish peroxidase.

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. I use 0.2-0.3 grams of BSA or lysozyme in 50 mLs of TBST. Using milk is fine but can impede detection of His6-tags.

(6) Disassemble the stack by reversing the assembly order. It's easier to peel the membrane off the gel. I immediately place my membrane in a plastic tip box lid containing about 10 mLs of blocking buffer. If you transferred a lot of protein and you don’t' put it in blocking buffer, the band will "flame" in the Western because unbound protein piled on the membrane will drift and stick to the membrane near the actual band.

(7) Pour off the first blocking solution and submerge the gel with about 25 mLs of fresh blocking buffer. Shake the gel in this for 30 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:    

(8) During blocking, unbound proteins wash off into the blocking buffer. So, pour off the blocking buffer and rinse once quickly with clean TBST.

(9) Add TBST with your primary antibody (I usually add a few mLs of leftover blocking buffer as well). The amount of antibody to use can vary greatly, but generally a 1/5000 to 1/20,000 dilution in TBST is plenty. Shake for 30-45 minutes.

(10) Pour off the primary antibody. Wash the membrane with 2-3 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. Remove an ECL kit from the fridge and open it. GE recommends having the reagents at room temp when used.

Light It Up!
(12) Decant the antibody solution. Wash the membrane well with TBST. I wash 4 times for 5 minutes and change the buffer immediately before adding the ECL reagent to get rid of any free enzyme. During the washes, turn on the film developer to allow it to warm up. Wipe the platform with a Kim-wipe sprayed with 75% EtOH to remove the rust specks that fall from the rusty ceiling vent. I usually cut my film now too so it's ready after the ECL soak. The film is made of plastic not paper.

(13) Mix the ECL reagent, decant the last TBST wash, pipette the ECL onto the blot making sure it's all covered. Tip the containers so the reagent drains to one corner and pipette it over the membrane a few times to make sure it is mixed with any remaining TBST. Incubate the blot with ECL 3-5 minutes. I re-pipette the solution a few times during the incubation to keep the membrane wet. For some reason, some people do this step in the dark room. The instructions don't say to do it in the dark. I do mine on my bench. They look fine.

(14) Lift the membrane out of the ECL reagent, let it drip for a second or two and place it face down on some Saran wrap. Fold the wrap up to cover the blot. Go into the Dark room. Generally, I place my blot on a piece of film for one minute to start. Close the cassette to press the blot flat against the film. Avoid shifting the blot when placing and removing it or it will streak your Western.


 * If you can see the bands glowing on the membrane in the dark room, one minute will be way too long. Just drop the blot onto the film for a few seconds and remove it.

(15) Feed the film in STRAIGHT to the developer. Use the platform edge as a guide along a straight edge of the film. Mis-feeding will cause the film to fall into the developer tank. It takes about 2:20 for the film to get through the developer. I place by blot on a second piece of film while the first one goes through.

(16) After the film drops out, look at it and decide how much longer (if any) the second exposure should go.

(17) That's about it.