Dear Peter

usually you need to take the maleimide in a 10-100-fold molar excess (e.g. as a solution in acetonitrile or other polar organic solvent). Which solvent to take depends on which organic solvent is tolerated by your protein. I think even trifluoroethanol should be possible as solvent. The reaction proceeds in 3-12 h in aqueous buffers. Hence, it should not be slower in an organic solvent. Separation from unreacted dye can then be performed using Sephadex LH-20 (the one that tolerates organic solvents) and gel permeation chromatography (a column of 20 cm and 2 cm diameter will be sufficient). Your labeled peptide will elute as a narrow band and the unreacted dye will follow in a broad weak zone. We did something similar a couple of years ago, but with a water-soluble protein (see attachment) where you need a different stationary phase.

I hope this will help you.

I had tried up to 30% DMSO/70%buffer that may help you. For our preps we have to incubate overnight at room temperature. You can also try the trick of dissolving your peptide first then, add some other miscible solvent. One issue that you are probably encountering with your peptide is the cross-linking of the Cys. Make sure to use DTT or TCEP to dissociate those before the reaction, remove the DTT quickly using a Hi-Trap column (size exclusion) and do add the maleimide-dye pretty quick. Some people keep a low concentration of EDTA (1mM) in the buffer to decrease the Cys to cross-link back. In our experiment we do not have to add the EDTA but something to consider. Another trick that we play is concentrating our sample down to 1 mg/ml solution before adding the dye. For our system concentration, excess of dye (100x) and DMSO concentration were all important. See for our experimental details at Soto C.M.,et al. (2008) Anal. Chem. 80, 5433-5440. Good luck.

Dear colleagues,

Thank you so much for your valuable suggestions!