Hi all!

I am trying to screen synthetic receptors for their ability to bind their ligand. The ligand has a His tag (Sino 50284-M07E), so I am using an anti-His Ab for flow (Biolegend 362637). I do a transient transfection, use ice cold DMEM to unadhere the cells and pass through a 70 um filter to de-clump cells. Then I incubate the ligand and cells at 4C for one hour, followed by staining with the antibody, and washing twice with FACS before flow. This did not work.

I did the following for troubleshooting:

• Changed to only using Cell Staining Buffer (BioLegend Cat. No. 420201) to unadhere the transfected cells, incubate with ligand, staining, and washing. I did this primarily because my protein binds calcium and I was worried the EDTA may have affected the protein conformation. I ran a native page to see if it did, and it did not appear to impact. For blocking, instead of having the DMEM, I used the TruStain FcX™ (anti-CD16/32, BioLegend Cat. No. 101319) and incubated on ice for 10 minutes. I did spin after this step and resuspended in cell staining buffer for staining.
• Tried crosslinking half the samples with 2% PFA for 5 minutes at 4C. As these receptors are surface receptors, I did not want to cause any permeabilization.

With both of these troubleshooting steps, I saw a significant increase in background. I am getting about 25% background with my blank cells + anti-His Ab, and up to 50% when my ligand is not present but the receptor is being expressed.

Is it too far fetched to use flow as a way to measure ligand-surface receptor interactions? I don't care about Kd or anything, I just want an easy way to screen if there is binding happening between receptor and ligand. Any advice on troubleshooting steps would be greatly appreciated! I am going to try titrating down the amount of Anti-his Ab for staining to try to decrease background, but thats the only next step I can think of.

Thanks so much!!