problem with Co-IP - (Sep/04/2014 )
hi,
Im using MC3T3 cells for Co-IP. I used the active motiff kit to do the nuclear extraction and used this for doing Co-IP. I precleard the lysate and did the usual protocol, washed with RIPA and ran the gel. The problem is Iam not able to see the IG bands in my sample and in the control sample. I used the SSB that I generally use for western and ran it on SDS-PAGE. Is this correct. Because I see protocols saying that beta mercaptoethanol should not be in the SSB and it can be run in native gel rather than SDS PAGE. Any suggestions are welcome.
Thanks
Jaya
I don't think that there is any problem in using the SDS PAGE sample loading buffer. I have used the same and I used to get bands for the light chains of the Antibody (Ab) on the blot, because we use light chain specific secondary Ab.
I don't think you are doing anything wrong by running your sample on SDS PAGE. By excluding beta mercapto ethanol you would get less of the heavy and light chains of the Ig on the blot, that seems to be the only logic. And anyways, the Ig bands are actually a nuisance on the blot.. :D But, I could not think of the reason why you don't get to see your Ab bands on the blot, which Ab you using by the way ?
In which species is your Ab raised that you are using for the Co IP and which secondary Ab you are using, may be that would be helpful and the only plausible explaination that I can think of.
P.S. My concern rather is that, is it common to use RIPA in Co IP ? I had the impression that it is better to use a milder detergent and not SDS atleast, unless you have really robust and strong interaction amongst your proteins of interest.
Good luck!
Thank you. The antibody I uses is all rabbit.
Hi jaya2020
Could you provide more information on your protocol and the specifics of your RIPA buffer? I agree with Poorti - this is somewhat uncommon to use in a Co-IP.
If the RIPA you're using contains SDS, you stand very little chance of having successful co-IP, since the SDS will readily denature your proteins by virtue of it being an ionic detergent. This greatly reduces the likelihood of any meaningful interactions being retained.
Co-IP is incredibly sensitive to subtle changes in the pH, detergent and salt concentrations of the IP and wash buffers in my experience.
If your goal is to see co-IP by denaturing/reducing SDS-page, then use the regular sample buffer to elute your sample. This is common practice. An alternative might be elution in low-pH primary amine (i.e. glycine) followed by neutralization with concentrated Tris ~pH 8.5.
Without more information it's hard to say what could be going wrong though. I'd suspect something is going wrong much earlier than the elution step if you see no IgG (washes, antibody concentration in the IP, western transfer, etc.) I presume you aren't using a secondary detection reagent that has low affinity to denatured IgG (Clean-Blot or True-blot, protein-A HRP, etc)?
Kind regards
I have don the Co-Ip in a nuclear extract which was prepared by using the kit from active mottif. They have given the lysis buffer in their kit and I dont know the composition of it. This extract is used for Co-Ip using the kit from thermofisher, where the antibody is immoblized to the resin. My result shows 2 bands which is seen in the test and in the control. So I thought there could be a problem using SDS PAGE as this is denaturing the proteins. Will a native gel be ok. Is there any procedure that can be modified.
I'm sorry but I'm having trouble understanding what you are referring to. What is your "test" and "control" in this experiment. What bands are you seeing (molecular weight/MW) on your gel? What are the MW of your expected target antigen, and the co-immunoprecipitated protein?
From the information you've provided, I'm guessing that you see two bands at 25kDa and 50kDa on your western blot. Is this correct? If so, you are visualizing the denatured heavy and light chains from your antibodies. These are omnipresent unless you use a secondary detection reagent designed to avoid cross-reacting with denatured IgG.
SDS PAGE is standard for analyzing IP. I don't understand why you want to use a native gel unless you are attempting to observe the co-IP'd complex itself. Even if that's the case, I think it is wiser to first validate your IP, then your co-IP by SDS-PAGE. Don't even bother blotting against your co-IP target unless you know you have a strong IP.
In general, what are all of the controls you're using for your experiment? For a co-IP, I would suggest the following controls:
1) Knockout/knockdown lysate IP'd with your antigen-specific IP antibody (Important if possible)
2) Nonspecific or isotype-copy IgG of the same species and isotype as IP antibody (critical)
3) Knockout/knockdown lysate IP'd with the nonspecific antibody (not critical, but helpful in noisy IPs)
4) Lysate Input lane containing a known % of your lysate used for IP (usually between 2.5-10% of the IP, critical)
I would suggest also carefully reading the details of your kits, and learning more about the general principles behind co-IP. It will make troubleshooting much easier - this is a notoriously difficult technique, so having more information will help you plan a more conclusive experiment.
Cheers
Thank you for the suggestion. My target protein is around 57kDa. i see one band around 57 and the other around 48. I want to used this result to study the protein -protein interaction by mass spec. The controls are the nuclear extract without the antibody and the test is the one in which the nuclear extract is bound to the antibody. The kit says the antibody is immobilised to the resin so the chances of elution with the final elutant are much less. since I want to use the final result for mass spec, I thought running on native gel would be better, that running it on SDS-PAGE. Am I correct in guessing.
I see - thanks for clarifying those points. So, to be clear, you're seeing these bands in both of your lanes? Do you mean you see them with one antibody, or two separate antibodies? Or when you say that you see two bands, do you mean that you see them on a stained gel?
In principle, if your IP is clean enough you can run mass spec on a 1D SDS-PAGE gel slice. This is easy if the proteins are close in molecular weight (otherwise you may need to cut 2, one at each MW). I've done this with pretty good success in the past. The native gel would have the advantage of allowing you to target both proteins in complex I suppose. So you could do it either way, I suppose. I wouldn't rule out SDS-PAGE while optimizing the IP though, since it will give you clear and predictable monomeric proteins from which you can draw conclusions about the efficiency of your co-IP.
Before any of that, though, I would be very careful and make sure you have a good assortment of negative controls. I can't stress this enough. It is bad practice to omit an IgG/nonspecific antibody control (and ideally a knockout control for one or both interacting partners).
From what I can tell, you only have a lysate (input) control - perhaps I'm mistaken? At a minimum, verify the specificity of your IP with an IgG/nonspecific antibody control before spending time and money on mass spec. This is very important because you have proteins migrating near where the heavy chain of IgG runs (55 kDa). IgG readily elutes in SDS sample buffer unless the antibodies are cross-linked to the resin (even then, I have seen some IgG elute with kits designed to prevent it). This is why many kits offer a low pH primary amine for elution, as opposed to a denaturing/reducing buffer.
Also, once you get to the mass spec point, I would encourage you to check out the Contaminant Repository for AP/MS experiments (crapome.org) to make sure that anything you do identify with mass-spec isn't merely an artifact of your solid-phase. They have a nice paper in Nature Methods: http://www.nature.com/nmeth/journal/v10/n8/abs/nmeth.2557.html
Cheers and good luck
Thank you very much. your suggestions are very helpful. Will try eliminate the background as you have mentioned.
jaya