Western blots may be great for visualizing protein expression, but they can be a perfect way to waste your precious antibody stocks if you follow the normal protocol. Thankfully, you don’t have to follow the normal protocol any more; here’s how to get great blots with a fraction of the antibody usage.

I have tried several different techniques to minimize the amount of primary antibody that I use.  In some labs, I have used heat-sealed bags to create a bag that is just slightly bigger than the blot.  This can significantly reduce the amount of liquid required to cover the blot.

More recently, without access to a heat-sealer I turned to diluting my antibody in the smallest volume of dilution buffer possible to just cover a blot laying in a Tupperware container that is slightly larger than the blot (usually 7-10mls for a 9×5.5 cm membrane).  Call me frugal, but even this approach still makes me cringe for antibodies that are diluted only 1:500-1:1000.

Floating your blot

But my newest (and now favorite approach) is one I call “floating your blot”. It requires a Tupperware container with a sealable lid and a flat bottom, parafilm, kimwipes, forceps, antibody dilution buffer and primary antibody.  A 9×5.5cm membrane requires only 1mL of antibody diluted into buffer.  To float your blot, follow these easy steps:

1)   Cut a piece of parafilm that is slightly larger than the membrane.

2)   Pipet some antibody dilution buffer without antibody (I usually use 1% low-fat dry milk, 0.1% Tween, PBS) onto the bottom of the Tupperware container and spread around using a kimwipe.

3)   Lay the parafilm onto the wetted area without creating bubbles between the parafilm and the container.  This sticks the parafilm to the bottom of the container.

4)   Wipe off excess dilution buffer

5)   Dilute antibody in antibody dilution buffer.  I use 1mL for a full membrane (9×5.5 cm), and 0.5mL for half a membrane.

6)   Pipette diluted antibody onto the parafilm near one end.

7)   Using forceps, pick up membrane that has been incubated in blocking buffer and lay protein side down onto the diluted antibody.  I lay one edge of the membrane against the liquid and allow the membrane to wick across. Keep the membrane on the parfilm and be careful to prevent any bubbles from forming between the liquid and the membrane.

8)   Seal container with lid.  This must be an air-tight seal or the diluted antibody will evaporate.

9)   Incubate with primary antibody using conditions optimized for the antibody.

10) Remove blot into new container for washes.

11) Technique can be repeated for incubation with secondary antibody if desired.

I wish I could take the credit for thinking of this on my own, but I didn’t.  A colleague showed me this trick and it really helps me save my antibodies.

What is the most useful tip you’ve received from a learned colleague?  We’d love to feature it in an article on Bitesize Bio – just drop us a line via the contact form to talk to us about it.

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  1. Hi Rebecca, thanks for sharing this tip. I tried it, and it gives clear bands. However, I have the issue of uneven bands (darker on both sides or one side of the blot), even in loading control (eg. B-actin). Is there a way to avoid this while using this format?

  2. Thanks for sharing Rebecca! I suggested this before, but it would be nice to see the real thing as for other “tips” or “gadgets” in the lab, maybe a visualized “experiment”?

    I fear the membrane getting slightly do the edge and some of it don’t get in contact with the antibody.


    1. Hi Alex, Thanks for the comment. As long as there is a centimeter or two of parfilm around the blot, I don’t think you need to worry about the membrane moving to the edge. I haven’t seen it move after any overnight incubation.


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