Chromatography Forum

Hello,

I'm woirking on an alkaline hydrolysis method for tryptophan. I take a powder with protein and expose it to NaOH 5N and cook in an oven overnight. Then I neutralize with 12 N HCL, centrifuge, filter and finally dilute.

My moblie phase is 50 mM phosphate buffer pH around 2.58 with small amount of TEA. An Agilent application chemist had me add KCl to adjust the ionic strength.

Peak shape: excellent
Peak Area Reproducibility: excellent

But tryptophan RT moves around so much. Is this a complicated interaction or chemistry

I did accidentally use the eluant described above to make serial dilutions of my standards....and I noticed it really helped but I'm concerned that if a use it with the unknowns the pH might destroy the tryptophan because the pH is pretty low.

I started getting very good recoveries when I made sure not to expose tryptophan to pHs below 7-ish. Oh, and when things got really wacky and I noticed that changing the guard helped bring on all fronts, but RT does still move around. The ghost peaks and system peaks seems to have consistent RTs but not tryptophan. My window can be 4.3 min to 5.3 min.

Thoughts anyone?

Can you share other information - column, flow rate, etc..?

I would personally use an acetate buffer mobile phase (ammonium acetate, 2%w/v) around pH 6.3 and also make sure the samples are at a final buffered pH of 6.3.

If you need to alter the retention time you can add ACN and reduce the aqueous mobile phase composition.

Your mobile phase is at a pH that's less than the estimated pKa1 of tryptophan (~2.83) while your sample preparations are kept around pH 7 you said?

There could be an issue with the tryptophan becoming protonated as the sample enters the system and comes in contact with the mobile phase, which could explain the shifts in retention time.

Unfortunately I don't have a ton of knowledge about protein HPLC analysis but this is all I can come up with.