Some attributes of large molecules make them behave differently from small molecules in reversed-phase separations.
Several readers have sent e-mail to me lately with questions and problems related to the liquid chromatographic separation
of proteins, peptides, and other large molecules, so I've combined their questions into a discussion of some of the problems
related to such separations. I'll use proteins as the model compounds, although the general behavior of large molecules is
similar. Reversed-phase liquid chromatography (LC) is the most popular separation technique for these molecules, and will
be the focus of this month's "LC Troubleshooting" installment. Reversed-phase LC, of course, is a denaturing technique, so
it is good for analysis, but not for purification or recovery of intact molecules. For example, if you desire to separate
an enzyme from other compounds and then collect it for other uses, you'll want to preserve the activity of the enzyme. This
means that the mobile phase must be sufficiently gentle to avoid denaturing the protein. Some techniques to accomplish this
include ion exchange, gel permeation chromatography (GPC), hydrophobic interaction chromatography (HIC), and affinity chromatography,
each of which uses a nondenaturing aqueous mobile phase. Reversed-phase mobile phases for protein separations usually include
acetonitrile, which will irreversibly denature the proteins.
When protein separations by reversed-phase LC were first being explored in the 1980s, some workers thought that the separation
mechanism was completely different than that for small-molecule separations. Today, we know that is not the case — the same
rules apply. But there are some aspects of the separation that we have to be careful of or we will not get the results we
expect. Even today these "surprises" can confound workers new to the field. Let's look at some of the aspects of large-molecule
separations that we need to pay attention to with reversed-phase LC.
Before starting any kind of LC separation, we need to pick a column. There are literally hundreds of reversed-phase columns
to choose from, but we still need to make a wise choice. With small molecules, typically we start with a silica-based column
with a C18 or C8 bonded phase. With large molecules, a C4 phase is a much more common choice. For small-molecule separations
(for example, <1000 Da), the sample molecules are small enough to get between the bonded phase chains on the packing surface,
so different chain lengths will effectively result in a different amount of chemically active surface area. Thus, we typically
observe that a C18 phase will have retention factors that are perhaps 70% larger than those for a C8 phase, or that it takes
approximately 5% more organic solvent to get the same retention time with a C18 phase as with a C8 phase. On the other hand,
large molecules (for example, >10 kDa) are too large to penetrate the densely bonded phase, so they only "see" the tips of
the bonded phase chains. This means that a C8, C18, and even C4 chain length "look" about the same to a large molecule. Another
way to think of this is to visualize the column packing as a toothbrush, where the handle of the brush is the silica support
and the bristles are the stationary phase molecules, fastened at one end. Small molecules might be thought of as grains of
sand that can penetrate into the densely packed bristles, whereas a large molecule might be more like a marble that sits on
top of the tips of the bristles. So there isn't much effect of bonded phase chain length on retention with large molecules.
Early in the development of such separations, a C4 chain length was chosen and has become the defacto standard for protein
A second aspect of the column that is important is the size of the pores in the packing particles. The silica support is not
a solid bead with the bonded phase on the outside, like the fuzz on a tennis ball. A better description would be a porous
sponge, or better yet, a popcorn ball, where the pores are the spaces between the kernels. As a result, nearly all the surface
area is within the particle, not on the outside. For this reason, the pores have to be large enough that the sample molecules
can easily penetrate the pores. As a rule of thumb, we want the pore diameter to be at least three times the hydrodynamic
diameter of the sample molecule. Columns used for small-molecule separations typically have pore diameters that range (between
products) of 6–15 nm (60–150 Å), which are ample for free movement of small molecules in and out of the pores. Such pores,
however, are too small for proteins, where pore diameters of 30–40 nm (300–400 Å) are more common. Larger-pore columns are
available for size exclusion separations, but surface area is roughly proportional to the pore diameter, and retention to
surface area, so excessive pore diameters may translate to insufficient retention.