You require 2 controls to measure IC50.

• MAX is the measurement when the inhibitor concentration is zero
• MIN is the measurement when there is no enzyme activity (this could be because you leave out the enzyme or because you add a concentration of an inhibitor that provides complete inhibition)

Once you have measured the activity of the enzyme (X) at each inhibitor concentration [I], calculate the % inhibition at each inhibitor concentration, as follows:

% inhibition = 100*(1-(X-MIN)/(MAX-MIN))

Finally, use a nonlinear regression program to fit the set of (inhibitor concentration, % inhibition) data to an equation such as the following:

% inhibition = 100*[I]n/(IC50n + [I]n),

where n is the Hill coefficient. Typically n is close to 1 if there is no cooperative binding of inhibitor molecules to the enzyme.

IC50 could be calculated using the following formula

IC50; 10^(log(A/B) × (50‒C)/(D‒C) + log(B), where A represented

the lowest concentration value at which the percentage

inhibition showed greater than 50%, B was the highest

concentration value at which the percentage inhibition

showed less than 50%, C was the percentage inhibition

value of the sample at concentrations B and D was the

percentage inhibition value of the sample at a concentration

A

Dear Mr Saphiro,

I have problem on determine IC50 on glucosidase assay. In my data the inhibition range only 5 - 40 %. In this case, is it still possible yo determine the IC50?

For this case, when determining IC50 using graphpad, should the data be normalized or not?

For inhibition that only goes to 40%, calculating an IC50 would involve an extrapolation, which is not advisable. You should increase the range of inhibitor concentrations to get more inhibition.

I don't know what you mean about normalizing the data. The IC50 is calculated based on % inhibition versus inhibitor concentration data. % inhibition data is derived from measuring the reaction rate in the presence of the inhibitor compared to the controls, as described in my previous answer.

following this question

Adam B Shapiro Thank you for responding sir, I have few more queries concerning the study. The protocol (attached) of the study says "The AMC standard is included so the researcher can quantitatively determine the specific activity of the enzyme using the AMC standard as a measure of how much AMC substrate was cleaved to release free AMC." and in the section "example of assay results" the protocol plots POP activity as POP (ng) v/s fluorscent intensity which is the plot of standard AMC. Now, how do I use this plot to determine enzyme activity and conclude which of my test drugs inhibited POP efficiently?

I gather from your description that this is a fluorescence assay using an essentially nonfluorescent substrate that is cleaved by the enzyme to release a fluorescent product, AMC (aminomethylcoumarin).

By preparing a set of solutions of known AMC concentration in the assay buffer (leaving out the enzyme), you can make a standard curve of fluorescence versus AMC concentration. This plot will be linear as long as the AMC concentration isn't too high. Since AMC is one of the products of the reaction and the two products are formed in equal amounts, the AMC concentration equals the product concentration. The slope of the standard curve is equal to the change in relative fluorescence units (RFU) per unit of product concentration (µM).

Using this slope, you can then convert the fluorescence measurements from your enzyme assay (measured under identical conditions as the standard curve) to product concentration (µM), as long as the fluorescence from the enzyme assay does not exceed the fluorescence of the standard curve.

The plot of POP enzyme versus fluorescence intensity you saw could be used to measure the POP enzyme concentration in an enzyme sample of unknown concentration, as long as the same enzyme preparation was used. This isn't very useful for most purposes. However, it is a good practice to generate a similar plot yourself to make sure the enzyme is behaving as it should. The plot should be linear as long as the enzyme concentration isn't too high.

All enzyme activity measurements should be made at the initial rate of reaction.

Adam B Shapiro, Sir, I did that and got a slope having R2 of 0.9954. From the equation of the slope, I have calculated the POP enzyme concentration of unknown concentrations. Is this slope only plotted for validation of the experimental protocol? The values of the calculated POP enzyme concentration (of unknown concentrations) can be useful for any calculations?

The information from that analysis is a measurement of the specific activity of the enzyme (micromoles/min/mg). This measurement can be used to compare the activities of different preparations of the enzyme, and can be used to measure the purity of the protein at each stage of a purification procedure.

If the substrate concentration was saturating, so that the enzyme was operating at Vmax, you can also then calculate the kinetic parameter kcat (turnover number), as long as you know the molecular weight of the enzyme and the enzyme is essentially pure.

Adam B Shapiro Please how can I select the working concentrations of a compund based on its IC50 value?

What do you want to do with the compound?

@Adam B Shapiro, I want to test its anti-inflammatory effect in vitro.

In a cell-free system, you can calculate the percent inhibition if you know the IC50 and the concentration because there is a simple, but nonlinear, relationship.

% inhibition = 100[L]n/(IC50n + [L]n) where [L] is the ligand concentration and n is the Hill slope, which is a measurement of cooperative binding.

If you are using a cellular system, you will have to do an experiment to measure the EC50 (50% effect) by varying the inhibitor concentration and measuring the anti-inflammatory effect.