We tried ion mapping by infusion for phosphorylation sites, and
found we missed things, because phosphopeptides are often low
stoichiometry, and often are quenched by presence of other peptides. For
phosphorylation sites, you need to have good coverage of the protein (just
because you've found one site, doesn't mean you have them all). This is
best achieved by LC/MS (I've seen phosphopeptides not stick to IMAC).
Generally this requires three LC/MS experiments. In the first two, we
analyze the total digest on oligo POROS resin, and the breakthrough from
the peptide trap on a high carbon loading column (to get the smaller
peptides). This gives you MS/MS of most of the peptides. We sort through
this looking for peptides that have undergone neutral loss of 98 or
peptides that have a mass that is 80 Da bigger than predicted (or some
multiple of 80). Often we find the phosphopeptide is a minor peak that did
not come up to the minimum in the original (they often trail in the LC),
thus the second LC/MS experiment is required to sequence that peptide
(along with other odd peaks we need more data on). In that case, its
usually a good idea to load more material; a good plan is to analyze 30% of
the digest in the first two LC/MS experiments, then load the remainder for
the second experiment.
If coverage is poor (many candidate phosphorylation sites are not
represented among the peptides observed), then you need to consider a
second digestion method. We like aspN. Beware of CNBr, because it can add
Br to peptides, which is 80 Da (although you can distinguish it from
phosphate by the isotope distribution).
Don't forget to be careful about spraying radiolabeled material.
If the sample has 32P, we let it sit in the freezer, until the
radioactivity has decayed. If we are doing outside samples, we check for
radioactivity ourselves (I've had a grad student say to me "it was only 200
cpm, so I didn't think it mattered", but that he had counted only 1% of the
sample).
Katheryn Resing
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