🚀 go-pugleaf

I have worked for some time with fluroescence polarization (FP)
in studies of drug-protein binding. My drug, which is
hydrophobic, typically has a fluorescent adduct and the
solution has 0.01% Tween 20 to keep down non-specific binding
to test tubes, multiwell plates, etc. Assay usually works well.

I usually do a check with a series of glycerol concentrations
to ensure that FP is working OK. I expect at high glycerol
concentration, the higher viscosity lowers molecular rotation
which is detected as an increase in the amount of polarized
light in the static (parallel) channel. This usually works
fine, fluorescein for instance. I recently have found some
labelled drugs that show no increase in polarization, and
even a small decrease (about 20 mP) even when adding glycerol
to 50%. Again, Tween 20 is 0.01% throughout. Fluorescence
intensity remains relatively constant with only a slight
parallel increase in both channels. The underlying physics of
FP always seemed pretty straightforward, but now I am
confronted with such data that has been reproduced several
times and it is not clear how this came about.

Strangely, when a protein that is known to be a drug target is
present, there is a strong increase in polarization (100-300 mP).
The glycerol data has created an uncertainty about negative
data for test proteins that I am screening for drug binding.

Is there anyone in here that might have some hints how this
lack of change in polarization in glycerol might have come
about, maybe some suggestions how to troubleshoot this?

Dominic