I'm visiting this forum for years, it is of great help.
Today, I have a persistent problem with my ChIP experiment. I will detail my purpose, protocol and hope you will be able to help me understand what is wrong. And I'm sorry for the long post!
I'm trying to study the recruitment on DNA of a coactivator (let call it X) for a nuclear hormone receptor (NHR). My model is the liver from mice either wild type or knock-out for the coactivator X specifically in the liver.
The coactivator activity of X over this NHR has been extensively characterized and the liver-specific KO validated through multiple approaches.
My ultimate experimental purpose is to realize the ChIP-sequencing of the coactivator X in WT mouse liver, and the one of the NHR in both WT and coactivator X liver-specific KO mice. But to be able to do that, I have to pass the ChIP experiment.
I perform the ChIP protocol on liver directly after its removal from the mice.
I've already optimized the PFA cross-linking and sonication steps.
FYI, I mince the liver in small pieces, wash it in PBS, fixe it for 10 min in 1% PFA solution (+ PMSF +DTT), quench PFA with glycine for 5 min, wash it thrice with ice-cold PBS, lyse with a dounce [...] and shear the chromatin using the Bioruptor Standard for 45 cycles (1 cycle = 30s "on" + 30s "off"), high intensity.
For the IP, I first pre-clear the chromatin using protein A-coated magnetic beads and incubating overnight (rotation, 4°C). Then, I pre-mix the coactivator X antibody with protein A-coated magnetic beads for 2-4h (+PI +BSA). As a IP negative control, I use a No Antibody condition. Then, I remove the beads from the chromatin and add the pre-mix antibody-magnetic beads. Incubation overnight (rotation, 4°C). Next morning, washes and decross-linking at 65°C overnight after adding salt. Finally, RNAse digestion, proteinase K digestion, DNA purification and qPCR using primers encompressing various validated genomic targets (or negative controls).
Now, my problems: in qPCR analysis, I don't obtain any significant and reproducible difference between the WT and the coactivator X KO samples.
Also, in many cases, I don't observe a significant difference between my "No antibody" IP and the ones using the Coactivator X specific antibody.
One thing that I hadn't test at first is the optimal quantity of the Coactivator X antibody per IP. I've just realized the experiment and you can see a qPCR results as a graph attached. In this experiment, there are two biological duplicates.
My analysis is that there is no significant differences between the negative control No Abs condition and any of the Coactivator X antibody ones. Second, there is no difference between the WT and the KO samples. The results are the same (same amplitude and scale) if I analyse a negative control genomic region. I analysed other positive and negative regions, same kind of results.
For now, I'm beginning to think that the antibody is not good. Or that my IP conditions are not optimal. Or else.
Why am I using this antibody? Because I've already performed the analysis of the coactivator X genomic recruitment in a different model (cell model) and that the results were significantly conclusive (and are almost published).
I've spent a lot of time trying to make this experiment works, and I'm running out of ideas.
I really hope that, with a fresh look, a clear and specialist mind, you will be able to help me.
Of course, I can provide any further detail that could be useful.
I will feel very thankful for any comment and advice!
I wish you a happy Holidays season,