In Part 1 of this series, we began our journey into the fascinating world of enzymology. We looked at the most basic concepts of what an enzyme is and the incredible jobs it can do. In Part 2 of “Working with Enzymes” I will look at some things to bear in mind, to keep your enzyme stable, purify it and measure its activity.

Keeping your enzyme stable

One of the most crucial things you must do when working with any enzyme is to keep it active for as long as possible. An inactive enzyme is no good for any experiment so understanding how to keep your enzyme stable under the assay conditions is vitally important.

So how do you keep your enzyme activity intact? Since proper folding is necessary for an active enzyme, most of your focus should be on preventing your enzyme from denaturing (i.e. unfolding). In the enzyme’s natural, intracellular environment, conditions are optimised so that the enzyme stays folded. However, when you strip the protective cell away and use the enzyme in the lab, it is much more vulnerable. Here are the main factors that you have to bear in mind:

Be cool

A folded protein relies on a plethora of intramolecular interactions to keep it together, including hydrogen bonds, electrostatic and polar forces. And according to the general principles of chemistry, the input of energy (often in the form of heat) can disrupt those interactions. Within living cells, conditions are such that the cellular machinery of chaperones, chaperonins, and protein-protein interactions keeps proteins folded- however, in vitro, these mechanisms no longer actively function to maintain protein conformation.  Therefore, you want to keep it on ice as much as possible. The more heat your enzyme encounters, the greater the chance your folded enzyme falls apart and loses activity.

Bitter and sour

The electrostatic interactions holding the protein together result from amino acid side chains that are protonated and deprotonated. A lysine must be protonated and positively charged (-NH₃⁺) in order to electrostatically interact with a deprotonated, negatively charged aspartate (-COO^-), for instance. Altering the pH of the environment surrounding an enzyme can result in excessive protonation/deprotonation, disrupting these sorts of interactions.

Salt in the wound

Salts can be a good thing. Sometimes they help improve the solubility of particular biomolecules – “salting in.” But add a bit too much salt and suddenly “salting in” becomes “salting out” and your enzyme precipitates right out of solution. Even before precipitation occurs though, an excess of ions will once again disrupt the necessary interactions between amino acid side chains that an enzyme needs to maintain its conformation, bind its substrates, and function properly.

Purifying your enzyme

Another big question that will ultimately arise is what level of purity you need in order to work with your enzyme. Can you break apart a cell culture, part of a plant, or piece of animal tissue and work with your enzyme right there in a crude situation? Or will you need to go through endless and diverse purification steps just to be able to use the enzyme to suit your needs?

Getting dirty

Whatever the source of your enzyme might be – a bacterial or eukaryotic cell culture, a plant sample, a piece of animal tissue – from the moment you begin your work you’ll need to take precautions to keep your enzyme functional. When cell membranes and organelle membranes are broken, enzymes and other biomolecules spill from their original compartments in the cell to make one big mixture, and you’ll need inhibitors to protect your enzyme from proteases and phosphatases that threaten to dephosphorylate or degrade it, potentially altering its activity and kinetics.

Up the long ladder and down the short column

For whatever reason, partial or whole purification of your enzyme may become necessary. How are you going to do this? In protein purification, the most common method of choice is typically some form of column chromatography. These days, automated methods such as fast protein liquid chromatography (FPLC) have become popular, where you can set up your FPLC and fraction collector with your column of choice and walk away from it. I personally still think there’s something to be said for more manual methods, like beakers, tubing, stir bars, and even Pasteur pipettes with long columns clamped to a lab bench, or inside a cold room- having your own hands do the work gives you greater control, which is especially important for those first few optimization steps. Particularly important during that optimization is figuring out what type of chromatography to use; are you better off using ion exchange, hydrophobic interaction, silica, or size exclusion? Usually a combination of these works best: look at the literature on similar proteins and see what other people did. If that is not an option, you may need to investigate different columns, but you will need an assay to see where your protein elutes…

Precipitation, salting, and concentrating

One of the possible steps you may encounter in purification is to precipitate either your enzyme of interest, or the contaminating proteins. This is when the previously mentioned “salting” methods come into play, using popular agents like ammonium sulfate to do the precipitating. An important question to ask is, “If you precipitate your enzyme, will you be able to resolubilize it and keep it active”?  Concentrating your enzyme may in fact be important following chromatography: if your enzyme ends up being spread across several elution fractions, the dilution will result in lower activity and require higher volumes to be assayed. Concentration may be necessary to squeeze more enzyme into lower volumes.

Keeping your enzyme active

We’ve been ignoring one of the main questions up to this point, haven’t we? How exactly are you going to assay your enzyme?

The ideal, perfect situation would be the following:

1)      You have an enzyme that has, for example, 1-2 substrates and 1-2 products.

2)      One of your substrates or one of your products is a chromophore that absorbs light – either UV or visible – at an easily detectable wavelength.

3)      The wavelength is one where there will be no interference from any of the other substrates/products, the buffers used, or – better yet – anything present in the crude extract/lysates of the enzyme’s source.

4)      You can use the “kinetic” mode of a spectrophotometer to detect the substrate while being consumed, or the product during formation.

Of course, often, this will not be the case. For one thing, just finding a biochemical reaction where substrates and products have different wavelengths can be difficult enough.

Cute couple

One solution that I’ve particularly enjoyed in enzyme assays is to couple my reaction of interest to a secondary reaction that consumes or produces a chromophore. Your primary reaction may not have any wavelength change, but your secondary reaction will. For example:

A + B  « Enzyme of interest » C + D          No wavelength change

C + X   « Coupling enzyme   » Y + Z          Z absorbs at 340 nm, nothing else does

However, it’s important to note that sometimes one coupling isn’t enough. One of the enzymes I worked with, creatine kinase, took not just one but two coupling steps to give me a chromophore I could work with. However, the substrates for the coupling enzyme, not to mention those commercial coupling enzymes themselves, got to be expensive.

Coming up…

We’ve lightly covered some material today, and a lot remains – especially if we want to give certain topics the in-depth attention they deserve. Next time, we’ll look at how to begin assaying our enzymes.