Thank you sir for your valuable comment. But actually I am looking for Urease Enzyme Inhibition assay. It would be a great help if you provide me one. Thank you again.
You begin with a urease activity assay. Then you add the inhibitor and measure the reduction in activity. You have to consider a few things.
1. How will you add the inhibitor? One common way to do it is to dissolve the inhibitor in dimethyl sulfoxide (DMSO) and prepare 2-fold serial dilutions in DMSO at 50 times higher concentration than the inhibitor will be in the assay. Add 2 µl of the inhibitor serial dilutions to the wells of a 96-well plate, then add 50 µl of 2X urea substrate followed by 50 µl of 2X enzyme. Mix with a plate shaker. Allow the reaction to proceed for a suitable amount of time. Add the detection reagent if necessary. Read the absorbance/fluorescence. You should include wells with DMSO but no inhibitor (MAX) and wells with no enzyme activity (MIN). For the MIN, you can use buffer instead of enzyme and DMSO instead of inhibitor.
2. What urea concentration should you use? A good practice is to use a concentration equal to the Km. That way, the assay is equally sensitive for finding competitive and uncompetitive inhibitors (The substrate concentration doesn't affect the sensitivity for finding noncompetitive inhibitors). You must be careful that the reaction rate is constant over the whole reaction time, which requires adjusting the enzyme concentration and reaction time.
3. Calculation of % inhibition and IC50. If your absorbance/fluorescence measurement is X, then % inhibition = 100(1-(X-MIN)/(MAX-MIN)). To calculate IC50, fit your % inhibition versus inhibitor concentration data by nonlinear regression to a suitable equation, such as the Hill equation: % inhibition = 100[I]n/(IC50 + [I]n) where [I] is the inhibitor concentration and n is the Hill coefficient.