Hi all,
We've been synthesizing some short (16 aa) peptides lately and they cleave beautifully from the resin, yielding a beautiful bulky white cloud in a 50:50 solution of hexanes/diethyl ether. Normally we further wash TFA out from them using this same solution over 3 rounds of centrifugation/resuspension of the peptide pellet, and further dry the precipitate overnight.
In the next day, we wake to see a white solid of (usually) dried peptides in our tubes, which prompt us to dissolve them into a 3:3 mixed solution of DMSO/Acetonitrile/miliQ water. The solution is completely clear as we inject it into the HPLC in a Water/Acetonitrile ramp gradient. Yet, each (high absorptive) tube will further yield precipitated crystals.
As we analyse the supernatant through LC/MS we see very few of our desired mass peak during the analysis, as we obviously never reconstitute the crystals in the injection solution (cause it could clog the system).
Would be interested to know what your thoughts are, and what you would do next.
Thanks!
Hi everyone,
For those who are following this question, I figured a way out of this problem:
When preparing my peptides for HPLC purification I usually follow a general protocol of dissolving them in a 1:1:1 ratio of DMSO/ACN/water.
Since crystallization requires a nucleation site, I believe dissolving peptides in the mixed solution I stated above can be problematic if you use MiliQ as your water source, as this may favor the formation of nucleation sites at given pH.
By replacing MiliQ by HPLC-grade water with 0.2% FA we favor solvation of the peptides and no more crystallization is observed.
Thank you all for following this thread and in particular Maksim Fomin for the suggestion!
Cheers!
Andre
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