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Get your stripping stripes! Find out how to strip and re-blot your Western

Westerns can be tricky and time-consuming, so make the most of your precious membranes and their proteins. Learn how to properly strip off your antibodies and re-probe with another primary antibody.

Why you should strip

Scientific reasons:

  1. To conserve protein samples that are limited or expensive.
  2. So that you can analyse the same sample with several different antibodies. This is particularly important if you want to compare relative abundance. For example: Comparing expression of protein isoforms, or measuring the levels of a phosphorylated protein in relation its total abundance. It’s also a useful technique if you want to look at two different proteins that have a similar molecular weight.
  3. To confirm your result with the same or a different antibody.
  4. To recoup your antibody for later use.

Real-life lab reasons:

  1. To remove an incorrect antibody (‘fess up – we’ve all done it!) or the right antibody at the wrong concentration.
  2. To save time and money by running less SDS-PAGE gels and transfers.
  3. To remove blocking buffer that has gone bad. This mainly applies to skim-milk blocking buffers that can go off at room temperature.

If you’ve decided that one of these reasons applies to you, next you need to learn how to strip. All stripping methods aim to remove your bound antibodies while leaving your proteins intact. There are three main stripping methods: 1) incubating with a low pH glycine solution, 2) using a combination of heat and detergent, or 3) washing in a commercial stripping buffer.

How to Strip

Method #1: Low pH

Commonly referred to as a “mild” stripping protocol, this method relies on a low pH glycine solution to dissociate bound antibodies. Low pH removes bound antibodies by altering their structure so that the binding site is no longer active. Low pH stripping buffers are made with glycine dissolved in water and buffered with a high molarity acid to a pH of 2.2. Sometimes 0.1–1% SDS (w/v) is also included to further help antibody dissociation. Typical incubation time for this buffer is 30-60 minutes. Expert tip: Don’t add SDS to your low pH stripping buffer if you want to collect your stripped antibodies for reuse. As we know from SDS-PAGE, SDS denatures proteins (like your antibody!) and antibodies only work in their native conformation! Also keep in mind that before you can reuse your reclaimed antibodies you need to adjust your stripping buffer to a neutral pH and allow your antibodies to re-adopt their native confirmation.

Method #2: Heat and Detergent

This is considered a harsher method of stripping but is appropriate for blots with higher signal. Like low pH stripping, this method works by altering the secondary structure of your antibodies releasing them from their target proteins. This is accomplished using a neutral Tris-HCl solution containing a reducing agents, such as beta-mercaptoethanol, and SDS. This solution is then heated with the blot at 50–80°C for up to 45 minutes with agitation. Expert tip: Unlike low pH stripping, recovery of your stripped antibody from this solution isn’t possible, as you can’t reverse the SDS and beta-mercaptoethanol denaturation. It’s also critical to wash your membrane really thoroughly after stripping with this method to ensure complete removal of the solution. Otherwise you risk your denaturing your re-probing antibody!

Method #3: Commercial Buffers (eg. ReBlot from Millipore, Restore from Pierce)

These buffers are all patented and so contain a bunch of mystery ingredients. Most claim to be gentler than old-school stripping methods — they don’t use mercaptans and don’t require high temperatures to work. They also strip your blot much faster than the traditional low pH or heat/detergent methods (most take only 15 minutes!) seemingly without sacrificing stripping efficiency. Commercial blots usually come in two strengths, covering your mild and strong stripping needs.

Think Before You Strip

Before you jump in and start stripping and re-probing, here are some important considerations:

  • Perform a negative control. The efficiency of your stripping protocol should always be tested before you re-probe. You can do this by adding developing substrate to your stripped blot and exposing it for detection as normal to confirm that no signal is generated. If you still see bands, you need to strip for longer or use a harsher stripping protocol.
  • Double check your proteins are okay. The effects of your stripping protocol on your bound proteins-of-interest should be tested. This is particularly important to test before stripping and re-probing for a quantitative analysis. If you fail to check whether or not you have stripped off your proteins, then your results can only be considered qualitative. So take the time to optimise your stripping protocol by comparing your blot’s signal strength (with a given antibody) both before and after stripping. If you see a drop in signal strength, you know that you have accidently also stripped off some of your target protein. If this is the case, use a gentler stripping method or perform shorter incubations.
  • Check that your detection method is compatible with stripping. Stripping only works if you have used a chemiluminescent or radioisotope detection system. As none of the above stripping methods will remove colorimetric substrates such as BCIP, 4CN, DAB and TNB. As these substrates form a physical precipitate on your membrane that is separate to the antibody itself, so dissociation of your antibodies won’t remove the coloured bands.
  • Keep your blot wet. Your blot can’t be allowed to dry out at any point after transfer as this will cause your antibodies to permanently bind to your membrane. To learn more about how to treat your western blot membrane read my other article Equilibrating your way to a perfect Western blot.
  • Remember to re-block! All of these methods cause dissociation of your blocking proteins as well as your antibodies, so if you don’t block you will end up with a black blot when you expose it. If you’ve have done this, don’t worry – just use your new stripping skills and start again!

The only question remaining when it comes to stripping is “how much is too much?” While I have never done more than 4 stripping and re-probing cycles myself, some researchers have successfully stripped membranes 10 times without any loss in signal. That’s a lot of time and sample saved! What about you? How could stripping improve your Westerns?

1 Comment

  1. Joe on June 10, 2016 at 1:59 pm

    Where can I find the recipe for the low-pH glycine stripping buffer? Thanks.

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