Enzyme assays

This is a bonus post to the one about enzymes; it's not a long one either. This time, I'm asking:

how do we know how effective an enzyme is at doing its job - either the rate of catalysis, or the change in concentration of a substrate - depending on certain conditions?

We work it out using an enzyme assay, which measures the conversion of substrate to product. Typically, we would measure this using the rate of appearance of the product, or the rate of disappearance of the substrate, both of which can be done using UV/vis spectroscopy - provided the substrate/product absorbs light at a wavelength within the UV/vis light spectrum. We could also use a fluorimeter to measure the change in fluorescence of the substrate/product to do essentially the same thing, again if they're fluorescent.

Some cuvettes used in UV/vis spectroscopy

We would also need to take into account other factors, such as temperature, pH, and any cofactors required for the enzyme to function. Additionally, you might also want to consider which solvent you're using in the assay, and the solution's ionic strength. These factors, along with several more, need to be considered to ensure the enzyme is operating at optimum conditions in your assay.

If the substrate/product is either not UV/vis-active or fluorescent, we can get around the issue by using an artificial substrate which is, effectively linking one enzyme reaction with another. Another approach would be to use two different enzymes:

• First, we'd have the non-UV/vis-active substrate converted into a product using an enzyme.
• Then, a different enzyme will convert the product into a different compound.
• For this to work, we need the second enzymatic reaction to not be rate-limiting, otherwise we wouldn't measure the effectiveness of the first enzyme. We can do this by having the second enzyme in excess of the first.

An example of a linked assay involves glucose oxidase and peroxidase. Glucose oxidase is an enzyme that catalyses the reaction of glucose with oxygen in the blood into gluconic acid and hydrogen peroxide. However, neither glucose nor gluconic acid are UV/vis-active; if we wanted to measure the glucose concentration in the blood, we'd have to link this reaction with an enzymatic reaction that does. Peroxidase does exactly that, using a cofactor to convert hydrogen peroxide into water, which brings about a colour change; have peroxidase in excess, and you can now measure the glucose concentration. This is especially useful in glucose monitoring strips for people with diabetes.

So long as our assays follow these conditions, we can use a continuous assay to determine the rate of conversion. Here, we'll record a series of points in one go with our spectrometer or fluorimeter, ending up with lovely graphs indicating our rates. These are great - they're not time-consuming, and we don't need multiple different samples. There is the notable caveat that using artificial substrates might not accurately mimic enzymatic behaviour within the body, along with various other issues outside the scope of this post, but otherwise it's great.

If our assays don't satisfy those conditions, we can use endpoint assays instead. The method is simple:

• Measure enzyme activity at certain times
• Repeat until you have multiple data points

This does mean you need way more samples than before, meaning the process takes up way more time and work than you'd want to do. That also makes the process more expensive, too.

When designing an assay, you'd also want your measurements to be reliable and reproducible. Considering for a moment the Michaelis-Menten equation that dictates rate of catalysis against substrate concentration, this means you want to focus on the region of linearity - where the rate is directly proportional to substrate concentration. The only issue with this is, depending on the enzyme, this region might be extremely narrow, and if you're going to do repeated measurements with frequent pippetting, that introduces significant human error and limited data. To solve this issue, we can have a large substrate to enzyme concentration ratio to make the region wider.

You might also want to make an assay to test the potency of an inhibitor; noting an inhibitor will affect the K_M and v_max values, you first need to establish what these values would be for no inhibitor. Now you'd add your inhibitor, and the rest of the process is the same. From this, you would then use a Lineweaver-Burk plot to determine what type of inhibitor is involved, which I wrote about in my enzymes post.