Moore:Chemiluminescent: Difference between revisions
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There are numerous sources available. Some solutions that are 33% are cheaper than 30%, so you can buy those and adjust the volumes accordingly. Get clean stuff and keep your fingers out of it. If this reagent goes bad, the system won't work. It is a good idea to have fresh aliquots used avery few months. | There are numerous sources available. Some solutions that are 33% are cheaper than 30%, so you can buy those and adjust the volumes accordingly. Get clean stuff and keep your fingers out of it. If this reagent goes bad, the system won't work. It is a good idea to have fresh aliquots used avery few months. | ||
=== | ===Luminescence Solution=== | ||
This was updated to make the mixing more straightforward. | |||
Pour ~2 mL of 500 mM Tris solution into a 15 mL conical tube. | |||
Squirt clean water into the tube to make 10 mLs at ~100 mM Tris. | |||
Add 50 μL of the Luminol stock (1.25 mM final, it will cloud briefly before you mix it into solution) | |||
Add 25 μL of Coumaric acid stock and mix (0.225 mM final, this can sit on the bench while you finish washing your membrane) | |||
Just before you pour off the last wash, add 3 μL of hydrogen peroxide and mix well, the reagent is now active. | |||
If you are unsure about the reagents, you can mix a sacrificial tube and pipette in ~0.1 μL of an HRP-conjugated antibody. The tube should glow in the dark for a few minutes. | |||
===Procedure=== | ===Procedure=== | ||
Revision as of 11:03, 18 December 2013
Chemiluminescent HRP Detection for Western Blotting
Protocol Background
This protocol is presented as a translation from a sheet of paper I received from a colleague. It describes a luminol-based chemiluminescent protocol for detecting horseradish peroxidase-conjugated antibodies during Western immunodetection. This protocol serves to replace the commercially available "ECL" and "ECL PLus" technologies sold and licensed by GE Healthcare. The Protocol I obtained is titled "ECL SOUTHERN / WESTERNS DETECTION REAGENTS" with the footnote "adapted from St. Geme and Miller lab protocols 1/4/01 by her highness Ms. J. Sexton". I don't know who these folks are, but thank you very much, you save us a lot of money.
Recipes of Stocks
250 mM Luminol
Buy Luminol (C8H7N3O2), and dissolve it in DMSO. CAS # 521-31-3 AKA: 3-Aminophthalhydrazide AND 5-Amino-2,3-dihydro-1,4-phthalazinedione Keep it in the dark when not in use (refrigerator is fine if your Fridge Penguin turns out the light when you close the door).
500 mM Tris-Cl, pH 8.5
If you need help with this, you shouldn't be doing Westerns. Diluting Tris stocks substantially will change the pH. Also, be mindful of the temperature when you pH a Tris solution.
90 mM ρ-coumaric acid
CAS # 501-98-4
Also dissolved in DMSO. Keep this dark as well.
30% hydrogen peroxide
There are numerous sources available. Some solutions that are 33% are cheaper than 30%, so you can buy those and adjust the volumes accordingly. Get clean stuff and keep your fingers out of it. If this reagent goes bad, the system won't work. It is a good idea to have fresh aliquots used avery few months.
Luminescence Solution
This was updated to make the mixing more straightforward.
Pour ~2 mL of 500 mM Tris solution into a 15 mL conical tube.
Squirt clean water into the tube to make 10 mLs at ~100 mM Tris.
Add 50 μL of the Luminol stock (1.25 mM final, it will cloud briefly before you mix it into solution)
Add 25 μL of Coumaric acid stock and mix (0.225 mM final, this can sit on the bench while you finish washing your membrane)
Just before you pour off the last wash, add 3 μL of hydrogen peroxide and mix well, the reagent is now active.
If you are unsure about the reagents, you can mix a sacrificial tube and pipette in ~0.1 μL of an HRP-conjugated antibody. The tube should glow in the dark for a few minutes.
Procedure
You will have more consistent results if you bring the solutions to room temperature before using, but it is not essential.
After your last wash of your "blot", mix the "Active Detection Solution" shown above. For mini-gels, making 1-1.5 mLs is plenty, just cut back on the volumes.
Drain the last wash off, do not let the membrane dry.
Pipette/pour the active solution onto the blot.
Mix it a few times by tilting the blot and pipetting the solution over the surface. Do NOT touch the membrane with your tip.
After a few minutes (I use 5 min, make sure you don't see areas drying), lift the membrane out of the solution either with tweezers or by sliding a pipette tip under the edge and grabbing a corner with gloves, and place it face-down on clear plastic wrap.
Fold the wrap to prevent the excess solution from leaking and expose to film in a dark room.
Expose according to experience. What this means is the signal intensity can vary greatly depending on (1) how much target protein you loaded on the gel, (2) How well presented the epitopes are on the membrane after transfer, (3) how specific/strong your antibody is, (4) how well conjugated your antibody is, (5) how old the reagents are... you get the idea.
I start with a 15 second exposure, then quickly move the blot to an unexposed area for a minute, then develop the film. Sometimes even that is too long of an exposure, cut back on something to weaken the signal the next time you do one.
Comments
Please, if you have used the protocol, leave some comments for others to help.
I don't understand why there is an additional dilution of Tris-Cl as a separate reagent, why not just make Detection solution #1 have 200 mM Tris-Cl? Has anyone cut "Detection Solution 2" out of the mix?
