Chromatography Forum

Hello,

I have to quantitate very low concentrations (<10 ug/mL) of a peptide (~4kD) and a protein (~55KD) in a solution.
Ideally one HPLC method would of been best, however...
The protein crashes out in the higher organic component of the mobile phase when using reverse phase chromatography.
So I have since developed an SEC method for the protein, and a RP method for the peptide using a Luna C8 (3 µm 100x2 mm 100 Å) column.

The Problem
I need to separate the peptide from the protein for this analysis. Using a centrifugal filtration method, the peptide aggregates to the protein, and nothing comes through. So, I was wondering if experimenting with varied amounts of SDS in the sample to prevent aggregation and then using an Amicon with a 10KD cutoff. And then using RP on the filtrate, for the peptide quantitation. But I am not sure of the highest concentration of SDS I can use without it causing damage/if any to the column. The column will be dedicated to this method.

I am not even sure if the SDS will prevent the aggregation of the peptide with the protein. I can only try. Not being a protein chemist, I feel I am doing this blindly. This whole project has been very challenging, I now have a deep respect for protein chemists.

One other thing, I have just read that there are these guard columns you can attach to your column that can remove SDS from your sample. Has anybody used these?

In the soap and detergents industry, we often injected solutions high in sodium dodecylsulfate and sodium dodecylbenzenesulfonate onto reverse phase columns with no issues.

We did use barium MaxiClean cartridges from Alletech occasionally to pull out fatty acids and soaps, but never explored those for synthetic detergents.

Our company sells guard cartridges that can be attached to reversed-phase columns. They're called "SDS removal cartridges". These remove SDS from samples but let peptides and proteins pass through. Once the acetonitrile level is > 70% (long after the peptides and proteins have eluted), the SDS elutes as well, preventing it from accumulating on the cartridge. These are available in i.d.'s of 1, 2 and 4 mm. If you expect a sample to have an unusually high concentration of SDS, then use the 2-cm cartridge instead of the 1-cm length.

An alternative would be to run your sample in the HILIC mode. The SDS would elute in the void volume. A decreasing ACN gradient would then elute first the peptide, then the protein. The only concern is whether or not the protein would remain soluble in the starting mobile phase. You can test this with a solvent consisting of 75% ACN containing 50 mM formic acid. It may be that these conditions will dissociate the peptide from the protein, in which case you wouldn't need SDS in the first place.

Thankyou Consumer Products Guy,
That is good to know.

Thankyou Andy,
It was your company's SDS removal guards I had read about. I will definitely give them a go.

I had briefly considered a hilic column, but having the protein precipitate recently whilst trying RP caused so many issues and interferred with quantitating the peptide. as I had random bits of the protein coming off everywhere.

In the end I had to wash the whole HPLC system out (minus the column) with hot water, phosphoric acid, an organic solvent, water etc...for a good 24 hours.

I may look at HILIC again, if we don't have to use SDS that would be bonus. BTW....Why the 50mM Formic acid?

Thankyou so much for your advice.

The 50 mM formic acid acts to solubilize the protein. Let's take the example of bovine serum albumin (BSA). If you put BSA into a solution containing 70% ACN (acetonitrile) and something like 10-20 mM overall of ammonium acetate or formate, then the BSA will precipitate. If instead of the salt you have 50 mM (overall concentration) formic acid present, then the BSA remains in solution. Chaotropes solubilize proteins and peptides that otherwise wouldn't go into solution. That's why people use urea or guanidinium hydrochloride as solubilizing agents. Unbuffered acids are chaotropes. Formic acid is a mild chaotrope, while TFA is a stronger one.

The problem with running your protein via reversed-phase is that proteins are substantially denatured by reversed-phase conditions. This exposes the numerous hydrophobic residues that normally are sequestered in the core in aqueous media. Their numerous interactions with the reversed-phase surface cause many proteins to elute late, in peaks 15 minutes wide, or sometimes not at all. HILIC is more forgiving in this regard, provided that you can keep the protein in solution in HILIC mobile phases. Recently Davy Guillarme and his group have published some nice papers on the use of TFA as the additive when performing HILIC of antibody fragments. Good solubility and nicely shaped peaks, at the expense of suppression of ionization in mass spectrometry (Guillarme's group isn't set up to perform top-down proteomics so it's not an issue for them).

Thankyou so much Andy.

I thought the addition of the formic acid was to help with the solubility, but now I understand how and what happens to the protein.

I will try a HILIC column, because if I could quantitate both the peptide and the protein in one method that would be great.

One of the other issues I found was since I needed sensitivity (due to the very low amounts), I lowered the concentration of the TFA in the mobile phase as I thought this caused the baseline to increase. I am concerned the 50 mM concentration of formic acid will do the same, as they both absorb in the same UV region.

However, I am still going to try it as the benefit of one HPLC method and no addition SDS or using an Amicon filter would be a bonus.

Actually TFA and formic acid do not absorb light in the same range. Like most carboxylic acids, formic acid has a reasonably high molar extinction coefficient at wavelengths below 230 nm, a range where you might want to monitor for proteins or peptides. The inductive effect of the fluorine atoms in TFA shifts its absorbance to the blue; it doesn't absorb light significantly above 205 nm or so. That's why it was adopted initially as an additive to the mobile phases for protein and peptide chromatography. The baseline does rise as you run a gradient to increasing ACN content in TFA-containing mobile phases, but if you're monitoring 220 nm, then much of that is probably from a change in the refractive index of the solvent.

TFA is fine if you're using absorbance detection; not fine if you're using mass spectrometry. That's because it forms ion pairs tenaciously with basic residues, suppressing their ionization in the mass spec.