If you can detect the peptoids with the detector connected to the HPLC, usually a diode array detector allowing UV detection, then you should easily be able to tell if they are degraded by proteases without using MS. For peptides, detection is usually done at ~214 nm (peptide bond absorbance). If there are aromatic groups, then you can use 260 nm.

It sounds like you plan to use reverse-phase columns. Personally, I would be concerned about clogging up the column with undigested proteases. However, the proteases will probably digest themselves during the incubation, so they will also be reduced to peptides.

If the protease concentrations are similar to the peptoid concentrations, you will see the peptide fragments of the proteases in your chromatograms, so blank runs of just the proteases without peptoids will be needed to identify the protease peaks.

Trypsin is specific for amino acids with basic side chains (lysine and arginine), so if these are not present in your peptoids, then there probably is no point in using trypsin.

Without knowing the size of the peptoids you are working with, my first suggestion is to separate them from the proteases used in your assay by employing physical methods such as UF, dialysis, and SEC. This will ensure a protein-free view of your chromatogram, whether obtained through MS or a UV-coupled LC configuration.

Secondly, the pore size of your resin is critical for direct applications. Above 200 A pore sizes are needed to screen proteins and peptides simultaneously. Lower pore sizes will cause fouling, high backpressure, and loss of resolution. Peptides will bind and elute generally earlier than proteins in columns holding pore size above 200 A, but this will be complicated when considering the hydrophobicity of the peptides and proteins if the ligand is C8 or C18 (C18 must be the choice for PMF monitoring btw). Some of the hydrophobic peptides retain much and may be co-eluted with proteins. However, you may try this application by blank subtraction of individual protease injections. Incubate the alcalase or trypsin or elastase without additional peptoids and inject to the C18 column with >200 A and record the sample as the sample blank. Further, identify autolysis peptides along with the intact protease if they are present. Then inject your peptoids before and after the digestion to prove they are resistant to cleavage and to discriminate them from protease-related peaks. Use generally accepted ACN-H202 binary gradients and TFA for UV, and formic acid for MS as MP additives.

To deplete enzymes, heating and precipitating them by centrifugation could be an option but I am not sure whether this can remove all proteases. Furthermore, autolysis peptides will remain in the solution. Therefore cannot be assumed as sufficiently versatile.

Good luck

If your interest is the peptoid fraction, HPLC-C18 is a very adequate method. I get results very satisfatory when I inject the incubation mixture directly on C18-column (acetonitrile gradient) without enzyme inactivation.. Enzyme samples must have high purity grade. By using synthetic peptides with aminoacid sequences suscetible to each enzyme you can monitoring peptide products through HPLC analysis. You must do the analyses of the incubation mixtures with active enzyme and another one with inactivated enzyme (heat inativation before the incubation - control tube). And then you can compare the HPLC profiles(peak retention time) by UV detection at 214 nm.