I’ve run into a problem and seek your advice.
I have done an IP but the problem is that two proteins I’m looking at, both have antibodies raised in the same animal, i.e rabbit.
and I get considerable backgroung on the blot after developing with ECL, such that it is difficult for me to identify my band of interest among so many.
Is there a way around to improve on that?
So, let me get this straight: you’re immunoprecipitating using a rabbit antibody, and you need to examine the immunoprecipitated proteins using another rabbit antibody?
I’ve no relevant practical experience, but it sounds to me like you’ll struggle to avoid background if you detect the second antibody using a secondary anti-rabbit antibody. Still, purely on a theoretical basis, a couple of options arise:
- I assume you’re immunoprecipitating using protein A agarose or something of the sort. If instead you were to covalently link the first rabbit antibody to beads, and rinse those beads really well before use, then elute your precipitate from the beads with a denaturant (leaving the first antibody still covalently bound), you might reduce your background enough.
- There are options for detection other than using a secondary anti-rabbit antibody. Back in the day you might iodinate the second antibody, but I think nowadays we’ve better, far safer options. Perhaps if you were to biotinylate the second antibody? Or to directly link it to peroxidase?
have you already tried reducing the amount of primary antibodies you are using or increasing the buffer stringency to get a better S/N ratio? This might at least give you a fighting chance of seeing the two bands that are important.
you got it right… my immunoprecipitating antibody as well as my immunodetecting antibody, both are raised in rabbit
and I am using anti-rabbit IgG as my secondary antibody on the western blot.
Could you send me the protocol you use for binding antibody to the beads (I use protein A/G beads from santacruz)
I will try reducing my primary antibody. I am right now using 5ug of primary antibody.
what is the protein concentration you people use for IP? I used 1mg (should I reduce the protein amount?)
Another thing, shall I pre-clear my secondary antibody with acetone power of wild type N2 worms? will that help in reducing background?
I get a ladder like profile after the western blot. all sorts of bands appear on the blot.
As I said, I don’t have personal experience doing these things, so I can’t give you a protocol I’ve used. Lots of kits are available.
I haven’t used the Protein A/G agarose beads from Santa Cruz, but generally there are a number of steps/controls/precautions that you could look at in your own protocol. Below, in the order they would appear in a general protocol are some suggestions to try out.
You could try pre-clearing the worm lysate with rabbit IgG.
You could use titrate the amount of protein against the 1o antibody as 1mg seems like a lot to me (try 200µg and see how this looks: remember the antibody concentration needs taking into consideration here, see 3. below).
You can reduce the amount of 1o antibody as 5µg is far too much (again this should be titrated, start with 1µg and see if that works).
What are you using to wash your immunoprecipitate?
I am using my HEPES lysis buffer for washing the composition goes like this
Triton-X 100 (0.1%)
Sodium flouride (50mM)
P.I before use
OK, well you could try uping the Triton X-100 a little, but not much as you will start to lose what’s bound, or try increasing the [salt].
Alternatively, if the beads are pre-blocked then you could add the blocker to your wash solution. Or try pre-washing with cold PBS a few times.
1.) I will up my Triton-x 100 to 0.25% as a higher concentration would again hinder IP like you suggested.
2.) will try using 500ug of protein lysate as against 1mg.
3. will preclear my secondary Ab with acetone powder of worms.
4.) increase washings of beads and add intermittent 2 washes with cold PBS.
5.) let the worms grow to confluency and will come back to you with the results.
or if I run into another problem for that matter.
As I understand it, and with the caveats I expressed above that I lack firsthand experience here, you still intend to elute the first antibody from the beads along with your immunoprecipitate and run it on the gel (the denaturant should disrupt protein A / protein G binding to that first antibody). As long as you’re doing this, and doing detection with secondary anti-rabbit, I don’t think you’re going to get rid of your background no matter how much blocking, washing, and diluting you do.
There are a few things that will help, but its hard to completely get rid of the background if you have the same species antibody for the IP and the detection in the western. I would recommend doing one thing from 1, and one from 2, and if your protein of interest is not close to the heavy or light chain bands, you may be ok.
- Leave as much antibody as you can on the bead and keep it out of your gel
a. As Hillel suggested, cross link your IP antibody to the beads.
b. Elute the IP from the beads with low pH glycine rather than boiling. Or if you have the peptide that your IP antibody is made against, elute the IP from the beads with that peptide.
- use a detection method in the western biased toward non-denatured antibodies (the primary in your western)
a. use anti-light chain secondary antibody for detection in the western (works the best for us).
b. Instead of an anti-rabbit secondary, try a Protein-A (rabbit) or Protein -G (mouse) secondary. These bind non-denatured antibodies best, so are more likely to pick up the primary antibody in your western, than what you used in the IP, which was then denatured and separated on the gel.
We’ve done these things with cell culture IP westerns and seen considerable improvement, but any of these might require a bit of titration or trial and error to find the best combination for your particular experiment. We generally find that we have to run through a few permutations with each new antibody we try.
not wanting to do this topic to death…but just some final observations.
Yes, you could pre-label with 35S but that too has its limitations and associated problems.
Yes, you could crosslink, but whilst it may improve your non-specific background it might also end up reducing your sensitivity. It’s also not without considerable extra work.
Yes, instead of boiling/denaturing you could try other elution methods to preserve the antibody structure, then it’s just a question of how well you separate the Ag from the Ab with such a elution buffer.
Yes, you could use another species…but that’s not without additional issues (costs, availability etc.).
Yes, (and this seems a possibility to me) you could use another detection system that avoids the issues of the secondary antibody. Something like HRP-conjugated protein A or G which binds preferentially to the Fc of the intact IgG.
Finally, the other suggestions I made, blocking, washing, diluting etc. were general approaches to removing/reducing background. As it is not absolutely clear from the first post in this epic saga whether the background (multiple bands) you see are ONLY related to LC/HC cross-reactivity, I thought it prudent to suggest controls to excludes other sources of background.
In any event, Renu, please let us know how you get on.
I get your point, that as long as I am doing it, I cannot get rid of the background completely but
even if there is a substantial reduction in the said non-specific bands I should be fine.
The band on the western should appear at around 75-80 kDa.
Thanks for your insights, using another detection system or another species,
well at present it is difficult to wait for these things given the time constrains I have.
In any which case, I’ll keep you guys posted on what did I finally get.
I think your suggestion of eluting IP complex with glycine to avoid excessive Ab transfer into the gel, seems to be quite doable
and might reduce the background as well.
I will surely try it.