Trying to quantify enzyme activity can be a messy business. In this post, I’ll discuss some of the complexities that can haunt both students and practitioners of enzymology. Since this post gets a bit discursive, all the meat is below the fold.
Usually enzymatic activity is quantified by adding a known quantity of an enyzme or enzyme preparation to some substrates, allowing an enzymatic reaction to commence. Substrate is consumed and product is formed. Substrate depletion — or, if it easier to measure, product formation — is measured over time, and the net rate of change of substrate or product concentration is usually taken as an enzyme activity.
Except, sometimes enzymes catalyze reactions which occur spontaneously under assay conditions, but at lower rates. An example is carbonic anhydrase. In one second, 3 or 4 out of every hundred CO2 molecules dissolved in water will spontaneously hydrate to form carbonic acid, H2CO3. Carbonic anhydrase carries out this same hydration, but it does it faster. If you happen to be tasked with measuring the activity of a given carbonic anhydrase preparation, what you need to do is measure the *difference* between the rate of reaction when enzyme is present and when it isn’t. Now, its the difference between reaction rates, not the absolute reaction rate with enzyme present, that is taken as the enzyme activity.
Confused yet? Don’t worry, it gets worse. Scientists have defined a veritable mess of different, related-but-distinct metrics for quantifying enzymatic activity. First there’s the unhelpfully named “unit”, (abbreviated “U”) of enzymatic activity. This is still the most widely used unit, even though the International Union of Biochemistry has been discouraging its use for at least the last 35 years. An enzyme unit is an amount of enzyme that will catalyze the conversion of one micromole of substrate to product per minute, at standard conditions. (More on standard conditions below.)
Because “minute” is not an SI unit, and probably also because “Unit” is a horrible name for a unit of measure, IUB defined the “katal” as the SI unit of enzyme activity. A katal is a catalyzed rate of one mole per *second*, not minute. (The katal, like the “unit”, is a measure of enzyme activity only in reference to a specified assay system or “standard condition”.) If you work through the arithmetic, one “unit” (U) of enzyme activity is equal to 16.67 nanokatals.
Enzymes are sensitive little molecules. Small changes in pH can lead to big changes in enzyme activity. The same is true for temperature. Substrate concentrations, product concentations, and even pressure, also affect enzymatic reaction rates. If one wants to compare enzyme activity from one experiment to the next, the conditions used to measure the activity must be standardized. Unfortunately, since the types of reactions catalyzed by enzymes are so diverse, no single set of assay conditions will work for all enzymes. Far from it, in fact. Instead, each type of enzyme will have its own “standard” assay system. If you’re lucky, your enzyme will have only one commonly used “standard” assay system. (Carbonic anhydrase, though, has had several “standard” assay systems in the past, leading to delightful appellations such as “Wilbur-Anderson Units” and “Roughton-Booth Units”, depending on whether you were using Dr. Wilbur’s preferred assay system from 1948 or Dr. Roughton’s from 1946. I told you it was going to get worse, and no, carbonic anhydrase is not alone in this regard.)
This business of “standard” assay conditions brings up two important questions. First, how can you determine what the standard conditions are for your enzyme? Second, what makes a good standard condition? Who designs these assays and how do they do it? The first question has a simple, but mostly unsatisfying answer: you have to mine the literature. If someone has studied your enzyme before, they probably had to create an assay for it. If multiple groups have studied your enzyme, they may have even agreed on the same standard assay conditions. And if your enzyme is a popular one, there may even be kits you can buy to do the assay.
If no one has ever studied your enzyme before, you are going to have to develop a new assay. And how will you pick the temperature? And the buffer to use? And the pH? And the concentration of substrates to use? The short answer: pick the conditions that maximize the rate of reaction. Pick the best pH for your enzyme; pick a temperature where activity is high but where the enzyme is stable. Perhaps most importantly, if possible, use substrate concentrations that are high enough to saturate the enzyme’s active sites.
The final question worth thinking about here is the connection between units of enzyme activity and the units of chemical kinetics. If we know the mechanism of an enzyme-catalyzed reaction, and we also know all associated rate constants, can we predict the activity that say one mole of enzyme molecules would have? The short answer is yes — *if* the substrates are saturating in the assay system. See the International Union of Biochemistry and Molecular Biology’s web page (section 12) for a short explanation — but as usual a more accurate answer is quite a bit more nuanced. But those nuances will be the subject for another time.