I want to reprobe my blot but don't want to strip unless I have to (so as to avoid losing protein - when I strip I use the pH2.5 glycine method which is supposed to be the preferred method for retaining protein but isn't necessarily the best for thoroughly removing antibodies).
I *know* I can use sodium azide to inactivate out HRP reporters and would prefer to do that whenever possible (I know to use this method you must have primary antibodies produced in different species) however I'm not sure what percentage sodium azide to use. I've seen 10% which seems unbelievably high. I was thinking 0.5-1% would probably do the trick but I was wondering if anyone knew for sure.
If anyone has experience or a recipe (especially if there's more to it than just dH2O and sodium azide) I would really appreciate it!
Hi, I have performed deactivation of HRP with sodium azide several times with 100% success. Here's is the procedure- prepare 100mM of sod. azide solution (0.6502 g NaN3 in 100 mL water). Add 100 ul in 10 ml of 5 % milk (blocking solution). Wash the membrane to be reprobed several times (minimum 3) with TBST/PBS. Incubate the membrane with blocking solution containing sod. azide. Best results are obtained with 8 hrs of incubation at 4 C. I kept such blots overnight in blocking solution. Next day, wash them 6 times with TBST (5 min each time). Probe with developing solution. You should not see any signal. Wash the membrane 3 times with TBST. Now your blot is ready for reprobing. Block the membrane again with 5% milk (this time without sod. azide). After 1 hr, incubate with new primary antibody.
Hi, I have performed deactivation of HRP with sodium azide several times with 100% success. Here's is the procedure- prepare 100mM of sod. azide solution (0.6502 g NaN3 in 100 mL water). Add 100 ul in 10 ml of 5 % milk (blocking solution). Wash the membrane to be reprobed several times (minimum 3) with TBST/PBS. Incubate the membrane with blocking solution containing sod. azide. Best results are obtained with 8 hrs of incubation at 4 C. I kept such blots overnight in blocking solution. Next day, wash them 6 times with TBST (5 min each time). Probe with developing solution. You should not see any signal. Wash the membrane 3 times with TBST. Now your blot is ready for reprobing. Block the membrane again with 5% milk (this time without sod. azide). After 1 hr, incubate with new primary antibody.
Some care must be taken in choosing secondary antibodies. If you use a rabbit anti mouse HRP for the first visualization and then block with the azide and then follow with an anti-rabbit HRP you will see the bands from the first visualization again due tot eh anti-rabbit rabbit-anti-mouse immunoreaction. If you have, for instance, a goat anti rabbit and rabbit anti mouse HRPs you want to incubate with the goat-anti-rabbit first.
The other issues is that the secondary will rarely react with 100% of the primary, so if you use, for example, rabbit anti mouse for the first protein of interest and then the same secondary for the next protein of interest you will see the first again... Simply saturating with secondary isn't the solution as you don't want too much signal when you visualize as you can get over-exposed blots that are useless for densitometry.
With proper planning a lot of these issues can be avoided. The first step is to test your antibody to see how specific it is and what concentrations are optimum. Once you know the band sizes you can plan the sequential incubations better to avoid having bands that overlap on the blot. For instance, I will perform the anti-rabbit incubations first, performing an azide block after each. With each incubation I see less of the first band I probed for until there is none left. Once I am done with the rabbit antibodies I move on to the mouse antibodies and work so that I am not going to get any overlap of the bands. I keep track of the sizes of the bands so I can exclude any band-overlap in the final analysis.
I would like to second Goodwin's warning about 0.2 M NaOH... The alkali will dissolve acidic proteins. The fixing step in acetic acid or TCA after blotting is performed to precipitate the proteins onto the membrane. They stay there unless exposed to heat, strong detergent (e.g. SDS) or alkali. As most stripping buffer use heat and detergents you will always lose protein with each strip and it is the more soluble proteins (e.g cytoskeletal proteins) that go first. And for some odd reason the stripping isn't uniform across the blot. If you perform any stripping you need to stain for total protein to be sure you are not losing your proteins of interest other wise you can end up with some strange results. I avoid striping at all costs.
Although this question is old, I will add my answer for anyone else trying to resolve the same problem. I have been doing streptavidin-HRP blots followed by HRP inactivation then probing with antibodies for protein of interest and loading control. Washing with 30% hydrogen peroxide, room temp. for 15 minutes worked really well for me as described in the linked paper.
https://www.ncbi.nlm.nih.gov/pubmed/19523435
http://www.sciencedirect.com/science/article/pii/S0003269709004011?via%3Dihub
I tried Nathaniel Jones' protocol (30% H2O2, 15 min, room temp) and it worked beautifully. Thank you :-)
@Hamida Safi
Did you use PVDF membrane based on Nathnaiel Jones' protocol?
Also, did it work for phorspho-proteins?
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