This is the first of a three part series describing some of the most common methods for concentrating proteins. In later installments I’ll discuss using protein precipitation and chromatography to concentrate a protein. However, here I’ll detail the most popular approach – semi-permeable membranes, used for both dialysis and commercial protein concentrators.
Structure of the membrane
Semi-permeable membranes are most often made of cellulose, and although they are diagramed as a solid sheet with little holes in it, electron micrographs reveal that they look more like a sponge. The size of the pores in the sponge-like structure determine the size of the molecules that can pass through the membrane. For simplicities sake, the manufactures translate this to a protein size, assuming that all proteins fold to a sphere-like shape.
This means there are a couple of things that you should bear in mind when using these membranes. First, the size and shape of the pores is irregular, so you should be careful when using a membrane to ensure that there is a significant difference between the size of your protein of interest and the rating of the membrane. Dialyzing your 11 kilodalton protein in a membrane rated at 10 kilodaltons will likely result in the slow disappearance of your protein over time. Second, your protein or molecule of interest may not be shaped like a sphere, allowing it to pass through the pores with some measurable efficiency. Assuming that your favorite protein is reasonably shaped (most are), and that you are able to purchase a product with a suitable pore size (several are available), then these membranes can be used to concentrate your protein.
Dialysis bags or cassettes
While dialysis is a great way to exchange the solution conditions of proteins or DNA, there is a property of the membranes that allows some control over the volume of the protein sample as well, and manipulating the volume of the protein solution is what protein concentration is all about. This special property? Water moves quickly and easily across the membrane, even faster than ions, which are ‘bulked-up’ with hydration layers. So when there is a strong difference in the solute concentration on opposite sides of a membrane, water will rush across the membrane to dilute the salty side more quickly than the salts can make the opposite journey. The result is that the volume of the original high-solute solution grows, and the original low-solute solution shrinks. This works against you if you are dialyzing a high-salt column fraction into a low-salt storage buffer, but you can make it work for you as well.
The principle that I described above for salt also works for glycerol. Many protein storage buffers contain glycerol to help stabilize the protein, but if you push the concentration up to the 40-50% range (as in many commercial restriction enzyme preparations), you can reduce the volume of your dialyzed protein fraction to as little as 20% of the original volume. This trick will even work if your original sample contains a much higher salt concentration than the dialysis buffer, although the volume reduction will be reduced to 30-50% of the original sample volume, depending on the salt concentrations.
This same principle is also exploited in many commercial dialysis-based concentration solutions. However, in this case large polymers are used rather than small molecules. These large polymers are, on average, too large to pass through the membrane pores themselves, which largely prevents the dialysis from coming to equilibrium. This means that the longer the dialysis bag/cassette sits in one of these solutions, the more water is extracted, and the smaller the volume of the protein solution. Consequently, much larger volume reductions are possible compared to glycerol solutions, but failure to properly monitor the dialysis could result in volume reduction beyond the critical point where the protein precipitates out of solution (or, if you’re lucky, forms a beautiful crystal).
If you don’t want to buy the proprietary mystery solution to concentrate your protein, you can do this yourself by using very high molecular weight preparations of polyethylene glycol (PEG) and a membrane with a very small average pore size. The PEG can be dissolved to form a very high concentration solution, or even used straight out of the bottle by laying a sample in a dialysis bag directly onto a bed of PEG chips, which will draw the water out of the sample.
One of the things to bear in mind when using either the commercial solution or the homemade variety is that the polymer solution will contain a mixed population of sizes, and that while most polymers adopt a spherical, globular shape in solution, they are not folded as a protein is and more readily adopt shapes that allow them to pass through the pores in the membrane and contaminate your protein sample. Therefore, this method probably shouldn’t be used unless the protein is going to undergo further purification steps that will remove possible polymer contaminants.
Commercial protein concentrators also use semi-permeable membranes, but substitute another force for the osmotic pressure that the dialysis methods use. The most common of these use centrifugation to force the protein solution through a membrane, but others use a pressure differential, either by pumping nitrogen into a sealed vessel above the protein solution or by creating a vacuum on the opposite side of the membrane from the protein solution. As with the polymer methods above, these concentrators have to be closely monitored to ensure that the protein sample doesn’t precipitate or completely run dry.
One common problem with these concentrators is that the rate at which the solution is passing through the membrane changes over the course of time. In other words, the first half of the solution passes through quickly, but then the volume appears to stabilize. Sometimes this is a sign of a disaster – your favorite protein is now embedded in the membrane, clogging it – but sometimes this is simply an effect of poor mixing of the sample. Before giving up on the sample or moving it to a new concentrator, try gently pipetting the remaining solution over the membrane several times (without touching it) and continue concentrating. To prevent this problem, many large-scale concentrators contain stirring mechanisms incorporated into the design. In addition, some centrifugation-based concentrators have also been specifically designed to avoid this situation, but you can negate this design feature if you use the wrong type of centrifuge rotor (fixed-angle versus swing-bucket), so make sure you read the instructions.
The Amazing Disappearing Protein
The biggest complaint about semi-permeable membranes is protein loss. This occurs for some proteins during dialysis, and for even more during force-induced protein concentration. It is an unfortunate fact of life that some proteins are just ‘sticky’ and love cellulose membranes. If you encounter this, and you’re your certain your protein didn’t simply pass through the membrane, there are a couple of things you can try.
Change membranes or manufacturers. Many of the common concentrators use regenerated cellulose membranes, but even though two different manufacturers both use this material to make their membranes, there will still be subtle chemical differences between them, and that might make all the difference in the world to your protein. Even within a particular manufacturer, there will be other types of membranes offered including derivatives of cellulose or polyethersulfone that you can try. Don’t be afraid to ask for free samples to test conditions.
Pre-treat the membrane. Another direction to try is to pre-bind likely binding sites on the membrane or apparatus with a (hopefully) inert protein like bovine serum albumin (BSA). I have personally had great success with this, but it comes at a price – you will have BSA contaminating your protein sample after dialysis or concentration, which could complicate determining the concentration of your protein of interest after the procedure. This also requires some forethought on controls, since you need to demonstrate that the contaminating BSA isn’t responsible for whatever effect you are attributing to your protein of interest.
Sometimes, despite all the tricks in the book, semi-permeable membranes just aren’t going to work for some proteins or for some applications. In parts two and three of this series, I’ll describe some other approaches to this problem.
This is part 1 of a 3 part series on the in’s and out’s of protein concentration:
How often have you looked at slides down the microscope and your thoughts have been miles away? Have you ever been sitting at the bench pipetting and preparing a PCR and wondered if you had really added your forward primer to all your samples (I’ll put my hand up to this one!)? Or spent time […]
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