I'm new to this forum.
I'm aiming to identify binding sites for a transcription factor. As no one has a good antibody against it, and in vivo expression is thought to be probably mainly embryonal, we use exogenous expression by transfecting with a V5-tagged construct instead.
I am trying qPCR with a few primer pairs: One in GAPDH promoter, two in macrosatellites, one of which contains the gene that encodes my TF, and the promoter and 3' UTR of a putative target for my TF.
Now, a control ChIP with H3-Acetylation antibodies pulls out the GAPDH promoter region quite efficiently, I get about 40 fold enrichment over IgG, or 2% of Input - I think the ChIP is probably working ok.
On my V5 antibody IPs, I get next to no signal in non-transfected cells for any of my primers tested (as I would expect). However, in my transfected cells I get a good enrichment for all my 5 primer pairs (about 10 fold compared to IgG, or about 0.5% of Input).
Because the difference between non-transfected and transfected cells is huge, I think it's not non-specific antibody binding, but rather non-specific protein binding. As I'm driving high levels of epxression from pcDNA3.1 with strong CMV promoter, I worry that I simply have so much protein that I just cross-link it to absolutely everything... Making my quest to find some binding targets (which we want to to by cloning of ChIPed DNA and eventually ChIP-Seq) impossible.
Any ideas? Why do you think I get enrichment everywhere? What could I do to solve this?
One of my main problems is that we do not have a definite positive control where we know my protein binds. But I worry that if I clone & sequence ChIPed DNA at this stage, all I'm going to pull out is a random pool of sequence that won't allow me to pick some nice targets.
Also, suggestions for a good negative control region are welcome.