In parts one and two of this series I described how semi-permeable membranes and precipitation methods could be used to concentrate your protein-of-interest, but there is one more method that you may not have thought of for protein concentration – chromatography. While chromatography resins are an obvious choice for protein purification, they can also be used to concentrate a dilute protein with a couple of modifications to the approach.
Unlike the other methods detailed in this series, this approach requires more knowledge of the protein of interest than just its molecular weight. Since we are often trying to concentrate a protein that we have already purified, however, that knowledge – what chromatography resins the protein will bind to and under what conditions it does so – is at our disposal. But, the protocol used with a particular column during purification of the protein may not be the best protocol for concentrating the already pure protein. Rather than trying to maximize the resolution power of these resins, the key to using them for protein concentration is to recover the protein from the column in the smallest volume possible. This means using the smallest column that will bind all of the protein in the sample and/or eluting all of the protein at once rather than allowing the protein to trail off a column. This mission is best accomplished with two types of chromatography resins – affinity and ion-exchange.
If your protein of interest was expressed with an affinity tag, most likely this tag was utilized in the first step of a multi-column purification because of their ability to remove the vast majority of the contaminating proteins in one step. However, if the tag isn’t removed in the course of the purification, then it can be quite handy for concentrating the protein at the end of the prep. Maltose binding protein (MBP), glutathione-s-transferase (GST), or histidine tags are the most useful for this purpose. The only other prerequisite for this method to work is that the protein has been bound to, and extensively washed on, one to two other columns in order to remove the elution molecules (maltose, glutathione, or imidazole, respectively) from the affinity tag.
The next key to this approach is to use the smallest column (volume of resin) possible to bind all the protein in your sample. Oversizing an affinity column generally has little effect on the purification power of the resin, with the exception of immobilized metal resins used for histidine-tagged proteins. However, larger columns lead to larger eluates and lower protein concentrations. Since you are now working with a purified protein, you can calculate the total amount of protein that you have, and better estimate how much resin you will need to bind it from the technical information that came with the resin. Due to the high binding capacity of these resins, you will likely be using less than a milliliter of resin, allowing you to collect very small fractions from the column, with most of the protein eluting in a single column volume. Alternatively, after washing you can load the resin into a disposable, empty spin column and elute the resin by centrifugation after a 5-10 minute incubation with the elution buffer.
Ion-exchange resins bind proteins via the specific ion moieties displayed on the surface of the matrix, and include Q (NH3+), S (SO42-), and Bio-Rex70 (COO–), amongst others. Unlike affinity resins, it is actually quite important to oversize an ion-exchange column during protein purification. This is because ion-exchange resins often don’t only resolve the protein of interest from contaminants during the binding phase, but also during the elution phase of the procedure. Consequently, you may or may not have access to small ion-exchange columns. If you are using a column in a FPLC, this may not matter, since these systems are designed to elute proteins in sharp, concentrated peaks. However, if you are using loose resin in a gravity column, then you will want to minimize the size of the column as discussed with the affinity resins.
When using an ion-exchange column for purification, the column is normally eluted with a salt gradient, and the ‘slower’ the gradient (ie – the rate that one moves from a low salt to a high salt concentration buffer) the better the resolution of the proteins, within reason. Sometimes under these conditions the protein of interest will elute in a nice sharp band, but some proteins aren’t as cooperative. When using an ion-exchange column for protein concentration, you can best ensure eluting your protein of interest in the smallest volume possible by not running a gradient at all, but rather to “bump” the protein off the column with a high salt buffer. The exact concentration of salt that you need is empirically determined from the purification profile, and I like to add about 50 mM to that just to be sure. For example, if the protein of interest eluted at 200 to 220 mM KCl during the prep, then I would bump the protein off the same resin with 270mM KCl when using the column to concentrate the protein.
One aspect of this approach that may be a problem for some proteins is that the protein of interest is now sitting in a buffer that is high in salt or that contains glutathione or maltose, requiring dialysis or a desalting (size exclusion) column to remove. However, this isn’t always a problem, as some proteins are best stored (ie – more stable) in high salt buffers, or the elution buffer may not affect the behavior of the protein.
Hopefully through this series I’ve given you some alternative approaches to consider the next time you need to concentrate your protein. If you have a favorite method that I’ve neglected to mention, please tell us about it in the comments.
This is part 1 of a 3 part series on the in’s and out’s of protein concentration:
Part 1: Semi-permeable membranes
Part 2: Protein precipitation
Part 3: Chromatography