I'm working with a coupled assay involving my main enzyme, Enolase, e two subsequent enzymes: pyruvate kinase, which transfer the enolase's product phosphate to an ADP molecule, producing ATP, and a luciferase, which produces luminescence proportionally to the amount of ATP. I'm sure about the assay conditions, since I've reproduced Enolase's Km (~ 30 uM).

Now, while testing inhibition, I'm having trouble. Some compounds were previously identified as enolase inhibitors, and I've tried to determine their IC50, without success (the enzyme worked basically as if without the 'inhibitor'). Then I've put a great quantity of one inhibitor (50 uM), and measured the amount of product after some periods of time. Usually I do the measurement after 20min, but this time I tried after 5min, 10min and 15min.

My readings were:

- blank: 71.986

- 5min: 71.644 (inhibitor), 98.738 (no inhibitor), 100% inhibition

- 10min: 114.952 (inhibitor), 223.576 (no inhibitor), 70% inhibition

- 15min: 178.138 (inhibitor), 347.565 (no inhibitor), 60% inhibition

I was expecting 100% inhibition, but got this result only measuring after 5min, when my '100% activity' reading weren't very far from blank reading, so I've not very much confidence about this result. Besides, my inhibition percentage variates with time, which I think shouldn't happen.

How can I improve my assay, so it allows me to identificate inhibitors and determine their IC50?