Yes, if your goal is to measure and compare the catalytic activity per molecule of the enzyme (turnover number or kcat) in different samples, then divide the catalytic activities by the molar quantity of the enzyme in each sample.

If your goal is to measure and compare the amounts of enzyme activity in different samples, then divide the catalytic activities by the total protein concentrations in each sample.

Adam B Shapiro

Hi Adam and thank you for your helpful answer.

I was a bit confused and hesitant to do the WB normalization, mainly because I was not sure about the role of my enzyme of interest in the background signal and non-specific signal (signals before correction).

Hi Adam,

In principle it does make sense to divide by the concentration of your enzyme of interest if your goal is to measure "specific activity" of an enzyme molecule rather than just "activity in a given volume".

However, the problem I think you will have is that Western blots are not quantitative enough for this purpose. You will probably end up with a lot of variation in your results that don't come from real differences between your enzyme activities but only from differences in your Western blots. A better method might be to express/purify the enzyme with His tags then measure the concentration by Abs 280 of concentrated samples prior to putting the enzyme in different conditions for activity measurements (a microvolume spectrophotometer can give an accurate reading from only 2 uL of sample as long as it is clean). Is this a possibility?

Good luck,

Jonathan

A more quantitative method than a Western blot is an ELISA. This requires a standard of the purified protein in order to determine the actual amounts of the protein-of-interest in the samples. For a relative quantitation (normalization), you could use an impure sample containing the protein to be used as a standard of comparison for the normalization. In either case, you would need to make standard curve by varying the amount of the standard and measuring the signal.

I'm having a little bit of difficulty understanding what you are after. If it's an average activity per mass of protein, you're ok but not if you're suggesting each molecule has lower efficiency. The expression system has an influence. As an example, when we were producing caspases in E. coli, if the expression was performed at 37°C, we obtained large masses of protein but very little activity. Expression at 15°C produced very small amounts of protein but with extremely high activity (10^3-10^4X higher). In the first case, the expression of the protein was so fast that it could not fold properly as it was being synthesized so we essentially had a large mass of denatured protein. I would be cautious about how you interpret the difference between your two conditions.

Dear Nick Chacos,

Thank you for your answer. I am mainly interested in the intracellular mechanisms that may regulate the activity of my enzymes of interest. I’d like to determine whether an enzyme is upregulated at the expression level but does not exhibit a corresponding increase in activity. This could be due to limitations in activation mechanisms (e.g. kinases that phosphorylate and activate the enzyme) or a concurrent upregulation of its inhibitory mechanisms (e.g. phosphatases).

So I guess I am thinking "each molecule has lower efficiency". But why does this make sense to me to normalize it using WB (or ELISA as suggested by Adam and Jonathan)?

ELISA makes sense since it essentially measures the number/concentration of target molecules regardless of whether there is any associated enzyme activity. You could also measure enzyme activity and get a ratio of active enzyme molecules/total enzyme molecules.