A) Do you agree this is the best stripping solution recipe :
Preparation of 100 ml Stripping Solutions (Final Concentration = 100mM):
2 gm SDS
6.25 ml 1M TRIS [PH=6.7]
Make it up-to 100 ml with water (Millipore)
Add 704.2 µL Beta-Mercaptoethanol
B) Please share the method you use for stripping the membrane i.e. incubation time period and condition
After stripping the membrane, I would like to reprob the membrane with another antibody. For e.g. Probing the membrane first with p-ERK1/2 and latter reprob with ERK1/2 primary antibody.
Thanks a lot for your expertise opinions.
Mild stripping
Buffer, 1 L
15 g glycine
1 g SDS
10 ml Tween 20
Volume up to 800ml with ultrapure water
Adjust pH to 2.2
Bring volume up to 1 L with ultrapure water.
Membrane incubation
Use a volume that will cover the membrane. Incubate at room temperature for 5-10 minutes.
Discard buffer.
5-10 minutes fresh stripping buffer.
Discard buffer.
10 minutes PBS
10 minutes PBS
5 minutes TBST
5 minutes TBST
Ready for blocking stage.
Harsh stripping
Prepare buffer and strip membranes under a fumehood.
Buffer, 0.1 L
20 ml SDS 10%
12.5 ml Tris HCl pH 6.8 0.5 M
67.5 ml ultra pure water
Add 0.8 ml ß-mercaptoethanol under the fumehood.
Procedure
Warm the buffer to 50°C.
Add the buffer to a small plastic box which has a tight lid. Use a volume that will cover the membrane.
Add the membrane. Incubate at 50°C for up to 45 minutes with some agitation.
Dispose of the solution as required for ß-mercaptoethanol based buffers.
Rinse the membrane under running water tap for 1-2 hours.
Traces of ß-mercaptoethanol will damage the antibodies. Wash extensively for 5 minutes in TBST.
You can buy stripping buffer from Thermo Scientific - we routinely use that rather than prepare our own.
http://www.piercenet.com/browse.cfm?fldID=02040210
make sure you wash your membrane a few times again before blocking again as you don't want the residual stripping buffer to react with the BSA or Skim Milk and ruin your blot!
0.5M NaOH for 15 ml at RT (25ml per membrane) on shaker. Followed by 2X washes in TBS-T at RT on shaker (5 mins ea) and 1X wash in TBS ( 5 min) at RT on shaker.
we also use the NaOH method for PVDF membranes; but i don't know if u can use it on NC membranes!
100mM glycine pH2.5 for 5-10min RT, twice.
then 2-3 washes with PBS or PBST, and block again with NFDM.
It's cheap, in house made, and you don't need to manipulate sds and b-mercapto.
hth.
best
Membrane stripping buffer: 100 mM glycine, pH 2.5, 200 mM NaCl, 0.1% Tween 20 (v/v), 0.1% (v/v) β-mercaptoethanol. Soak the membranes for 45 minutes at room temperature and then wash extensively in TBS. After repeating the blocking step, the blot can be re-probed. Reference: Derkinderen, P., et al. (2001) Dual role of Fyn in the regulation of FAK+6,7 by cannabinoids in the hippocampus. J. Biol. Chem. 276(41):38289-38296
100 mM Glycine, pH 2.0 5 min., wash with TBST three times, block 20min. 2.5% milk iin TBST
It depends what you would like to do later: For re-use of antibodies --> use the suggested Gylcine/HCl-buffer, pH 2.5. That's the same you are using for elution from affinity chromatography columns (but without Tween). This can be easily neutralized by a 0.1 mol/L carbonate buffer pH 9.6.
If you are adding ß-mercaptoethanol --> the antibody will be destroyed.
You can elute by using this method, band-specific antibodies from a polyclonal antiserum and see what the antibodies recognize else.
Also 3 M urea is fine, but needs several washing steps.
Don't use NaOH, this will destroy the antibody as well as parts of the antigen on the NC/PVDF-membrane. You will lose sensitivity.
Oh, OK... but I agree with Stephan... not the best idea...
Usually we do not reuse antibodies... but anyway mercapt stinks lie hell, especially when heated.
I also used 5mM glycine 150mM NaCl pH 2.4, but the one I write above works equally well...
Preparation of 100 ml Stripping Solutions (Final Concentration = 100mM):
2 gm SDS
6.25 ml 1M TRIS [PH=6.7]
Make it up-to 100 ml with water (Millipore)
Add 0.78 mL Beta-Mercaptoethanol, adjust pH to 6.7.
Procedure:
1. Warm the water bath to at least 50 C maximum 70 C.
2. Insert the membrane in the warmed stripping buffer present in a heat-resistance plastic box for 30 min (mix the buffer every 10 min). WARNING! Do the stripping in a FUNNEL as beta-mercaptoethanol is highly toxic!
3. Dispose the stripping buffer, fill the box with TBST for 30 min, change TBST every 10 min.
4. Control the efficacy of stripping with ECL/photo or alternative method to detect second antibody.
5. Repeat the procedure if needed.
Got this one from my PI at BUMC.
0.2M glycine (HCl) pH 2.0.
-Wash blots twice with 10 ml 0.2M (pH 2.0) glycine for 30 seconds.
-Then incubate blots for 20 minutes at RT in 20 ml 0.2M glycine (pH2.0) with gentle rocking. Incubating at RT works but 37oC is better.
-Then wash blots twice for 2 minutes in TBS-T buffer before re-blocking with 5% BSA (fraction V) or 5% low fat milk etc.
cheers,
Ewan
Incubate the membrane in the mentioned buffer at 50°C for 30 minutes with occasional agitation.
I have consistently used 1% SDS in dH2O, agitating at room temperature for 1.5-3 hours, depending on the signal (bound ab).
I agree that "the best" method depends quite a lot on what you're going to do with the membrane after. I would caution that the glycine method may not remove all of the antibody, it varies from antibody to antibody (I think mostly due to avidity). The first time you use it, you should re-probe with the secondary antibody again to be sure all the primary is stripped. When we are probing for two proteins (i.e. an experimental and a house-keeping protein), that are well-separated on the membrane, and if there is no cross-talk between antibodies, we just don't bother to strip off the first antibody. But again, you have to be cautious that both antibodies are specific before doing that.
we use this one:
Stripping Buffer 1.56 ml 1M Tris, pH 6.7 (62.5 mM final)
2.5 ml 20% SDS (2% final)
25 ul 2-beta-mercaptoethanol (100 mM final)
up to 25 ml with MQ water (make fresh)
My suggestion is that below mentioned changes are required in your composition-
Add 2gm SDS, 12.5ml 1M Tris Cl (pH 6.8). and after making volume to 100ml with double distilled water, add 700ul beta-mercaptoethanlo (Invitrogen).
Hi, I would think your method is ok, but it is quite strong and harsh stripping your antibody away. The Glycine method mentioned by Ewan Dunn is not as strong. So, I always go with such a acid Glycine solution instead of any 2-ME method. Keep in mind that with each stripping of your membrane you are loosing roughly also 50% of your protein of interest. Keep this in mind for the choise of your order for protein detection: proteins which are complicated to detect should be developed first and at the end loading controls liek beta-tubulin or so. Good luck!
I generally use Nitrocellulose membrane
-Wash blots twice with 10 ml 0.2M (pH 2.0) glycine for 30 seconds.
-Then incubate blots for 20 minutes at RT in 20 ml 0.2M glycine (pH2.0) with gentle rocking. Incubating at RT works but 37oC is better.
-Then wash blots twice for 2 minutes in TBS-T buffer before re-blocking with 5% BSA (fraction V) or 5% low fat milk
I use the same recipe for pvdf membrane during 30min at 55°C
then you have to wash every 10min during 1h with TBST
and re-block with 5% low fat milk
I'm always wondering what denaturing agents are used here. SDS denatures antibodies permanently (due to strong hydrophobic interactions with the protein [here the antibody]), ß-mercaptoethanol will brake the disulphide bridges of the antibody, NaOH will cleave/hydrolyse the Amino acid chain, heat > 40 °C will denature any protein, ...
If you would re-use either the antibody or the antigens on the membrane, there is only high molar urea (until saturation) or low pH thin buffers.
High molar urea needs some dialysis before the antibody can be re-used. If this is not required, this method is very effective and needs only several washing steps the the the urea removed.
Acid buffers (pH 2 ... 2.5) are also fine, need sometimes two or even three incubations. It has to be a thin buffer to get this easily neutralized. This has to happen within a short time to avoid hydrolysis of the antigen or/and the antibody.
If you need further information, look for basic principles in affinity chromatography.
Proteins in your gel are already denaturated, you can't denaturate it more !
Good stripping should removes the antibodies, but leaves transferred protein.
In my experience, good stripping depend on the antibodies and lysate. Some antibodies are stronger binding or the targets are more abundant. These antibodies are more difficult to remove. Start with the antibody that can be stripped the easiest and then follow with stronger antibodies.
Also, PVDF is more robust physically and chemically than nitrocellulose. It can withstand harsh stripping buffers and more stripping/reprobing cycles if required.
I've tried SDS/Tris/2-mercaptoethanol, Pierce's stripping buffer and guanidine hydrochloride buffer (from Yeung and Stanley, 2009, Anal. Biochem.) and have had successes and failures with each. Nothing is perfect.
Dear Laurent,
there is is difference in denaturation by SDS. This will leave the chain of amino acids intact. Some epitopes (linear) are still recognizable, others not, especially the one which depending on the conformation. If you incubate your proteins in acid or basic conditions, you will have a hydrolysis. This can destroy even linear epitopes. That's the reason why in affinity chromatography thin acid buffers (Glyin/HCl) are in use. They can be neutralized easily.
Heating a protein will leave undefined coils. A couple of epitopes are lost.
Especially when you are incubation the antibody containing solutions with SDS, ß-ME, NaOH, ... you could not do any further experiments with them.
What you are trying to do is to get the binding between antigen (on the solid phase PVDF or NC) and the antibody released. Therefore you need a change in the conformation of the antigen-binding site of your antibody (not a denaturation). This can be done either by acid buffers or high salt concentrations
1. Glcine/HCl
2. 8 mol/L Urea
3. 3 mol/L KSCN
4. Guanidine-HCl.
Mild stripping
Buffer, 1 L
15 g glycine
1 g SDS
10 ml Tween 20
Volume up to 800ml with ultrapure water
Adjust pH to 2.2
Bring volume up to 1 L with ultrapure water.
Membrane incubation
Use a volume that will cover the membrane. Incubate at room temperature for 5-10 minutes.
Discard buffer.
5-10 minutes fresh stripping buffer.
Discard buffer.
10 minutes PBS
10 minutes PBS
5 minutes TBST
5 minutes TBST
Ready for blocking stage.
Harsh stripping
Prepare buffer and strip membranes under a fumehood.
Buffer, 0.1 L
20 ml SDS 10%
12.5 ml Tris HCl pH 6.8 0.5 M
67.5 ml ultra pure water
Add 0.8 ml ß-mercaptoethanol under the fumehood.
Procedure
Warm the buffer to 50°C.
Add the buffer to a small plastic box which has a tight lid. Use a volume that will cover the membrane.
Add the membrane. Incubate at 50°C for up to 45 minutes with some agitation.
Dispose of the solution as required for ß-mercaptoethanol based buffers.
Rinse the membrane under running water tap for 1-2 hours.
Traces of ß-mercaptoethanol will damage the antibodies. Wash extensively for 5 minutes in TBST.
I agree with Sreedhar, in our lab we also use mild stripping buffer
15 g glycine
1 g SDS
10 ml Tween 20
Volume up to 800ml with millipore ultrapure water
Adjust pH to 2.2
Bring volume up to 1 L with millipore ultrapure water.
Incubate the membrane for 15 min in the stripping buffer
Remove the buffer and add fresh buffer again incubate the membrane in stripping buffer
Wash the membrane with 1xPBS for 1o min
Wash the membrane with 1xTBST for 5 min two times
The membrane is now ready for down stream applications again
You may use hybridization oven at 50°C. It helps to reduce exposure to beta-merkaptoethanol.
There is some disadvantage of the stripping buffer which contains SDS and b-Mercaptoethanol. It reduces phosphorylation of proteins, so I recommend to expose the mebrane with anti-phosphoAb to ECL for longer time periods (30-40 minutes).
Ultra pure water for stripping buffer?????
I agree glycine 0.2m ph2.2
1%sds. ,triton
Beta Mercator is fine
10min. Rt
Better to saturate the mb before reprobing
We used this method in our lab and it worked really well!!
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2679975/
Hi!
I gave up the mercapto buffer, because the whole lab was stinking days after the stripping and I was not the most liked person in the lab!
I tried for a while 50 mM NaOH solution for 1 hour. It is good, but only if your antibodies do not bind to strong.
At the moment, using a PVDF membrane, I am able to strip the membrane up to 6 times, without loosing signal of proteins. The trick is to let the membrane dry after blotting transfer (ONLY WITH PVDF), and reactivating shortly with 5 ml Methanol before blocking. For the stripping, I do at the moment: 25 mM Glycine, pH2 and 1% SDS. Since I am lazy by nature, I have done the stripping after luminol removal with water, then 30 min with the stripping solution, remove excess of stripping solution with TBS-Tween buffer and start the blocking again. VERY IMPORTANT: Do not let your membrane dry after the developing. If you do, your antibody will bind to the membrane and you will not be able to remove it at all.
I have made a 10X stock concentration of the buffer, so that nothing grows in it and that I can make new solutions "fresh" shortly before I need them. Only problem was in winter: Our lab got so cold, that the SDS precipitated, but 1 hour at 37°C and I could re-disolve it again.
Good luck!
Luisa
I have the same question too. Does anyone have the experience to prepare the stripping buffer in house?
We use the same procedure as you mentioned... i seal the membrane with the buffer and rock it in water bath for 20 min.. and wash with TBST *6 times 10 min each....but this is really harsh stripping sometimes i dont even see the signal....really faint...
Nandaraj Taye: you mentioned "5mM EDTA and boil the membrane in microwave oven for three minutes". can u tell me till what temperature you boil..can i do it at 55 degrees?
Hi everyone,
I use Abcam's protocol for mild stripping (with minor changes) and it works really well for me:
Mild stripping Buffer, 1 liter
- 15 g glycine
- 1 g SDS
- 10 ml Tween20
- Adjust pH to 2.2
- Bring volume up to 1 L with ultrapure water.
Membrane incubation
- Use a volume that will cover the membrane. Incubate at room temperature for 5-10 minutes. Discard buffer.
- 5-10 minutes fresh stripping buffer.
- Discard buffer.
- 10 minutes PBS
- 10 minutes PBS
- 5 minutes TBST
- 5 minutes TBST
- Ready for blocking stage.
I incubate for 30 minutes instead of 5-10m (1h total) and use PBST instead of TBST.
I have tried this protocol with several different antibodies, and I always get total stripping of the membrane, with negligible loss of signal. I am using PVDF membranes.
Good luck!
@Thao Tran..I adjust the pH first and then bring up the volume. The only problem you might face if you do another way is that if you need more acid/base to adjust the desired pH it may result in the increase in final total volume.
I took the one from Millipore guide to Western blotting:
25 mM Glycine-HCl, 1% (w/v) SDS pH 2
After detecting with peroxidases, I removed the reagent with water and add 5 ml of stripping solution for 1h under rotation.
I have been using PVDF membranes and from each membrane I can re-strip up to 6 times, enough to keep me a whole week busy. We have 2 antibodies that are extermely strong. I have tried to remove these completly but there is always something left, so I leave these ones for the last western blot of the series.
As an accident, I have left it overnight and even over the weekend: Overnight caused no damage. Over the weekend, let's say that the 6 re-stripping steps were reduced to 3 cycles ;-)
Mercapto containing buffer has caused more than one fight in our lab... I will really recommend to avoid it. It is not that good either (in our experience).
I tried the 5mM EDTA in the microwave for 3 minutes.... I wish ResearchGate allowed for pictures because it melted the container (no leaking) and dried out the western when the liquid evaporated.... I could see through the center of my PVDF membrane.... I guess mission accomplished, no more antibody. (Already had tried the mild and harsh stripping buffer on the blot, I was just trying to see if anything was stronger). I was told that getting the 5mM solution to near boiling, then dumping it onto membrane for 30 minutes shaking also works and is a bit less harsh. I will try that next.
I am using the following protocol
Abcam's Stripping buffer (pH 2.2) for 1 Liter
Glycine : 15 G
SDS : 1 G
Tween 20 : 10 mL
Adjust pH to 2.2
Bring the volume to 1 Liter with ultra pure distilled water.
PROCEDURE:
1. 10 MIN STRIPPING BUFFER INCUBATION
2. DISCARD IT
3. 10 MIN STRIPPING BUFFER INCUBATION
4. DISCARD IT
5. 10 MIN PBS WASHING
6. DISCARD IT
7. 10 MIN PBS WASHING
8. DISCARD IT
9. 5 MIN TBS-T WASHING
10. DISCARD IT
11. 5 MIN TBS-T WASHING
12. DISCARD IT
13. BLOCKING WITH 5% NON FAT MILK FOR 1 HOUR
14. 5 MIN PBS WASH
15. INCUBATE 1ST AB
THIS WORK VERY SUCCESSFULLY FOR ME. I STRONGLY RECOMMENDED THIS PROTOCOL TO EVERYONE. I USING THIS 3 TIMES
THANK YOU
KARTHIK
I have used Abcam's protocol for mild stripping.... it didn't worked for me at all.....
i got the same bands of previous antibody
http://www.biolchem.ucla.edu/labs/grunstein/Western%20Membrane%20Stripping.html
We use similar protocol with fewl changes for stripping PVDF membranes and it works fine.
Abcam recommends using a PVDF membrane but... does anybody know if their stripping protocol works also for nitrocellulose membranes?
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