I have a question about a problem I have when preparing samples for SDS gel electrophoresis. Usually I add 2% - 5% BME (or DTT) to denature disulfide bonds and heat the samples for 10 minutes. This usually works great and denatures the sample which then runs as a monomer. However, when I do this with mammalian transferrin receptor it seems to get dimerized or agggregates and runs at what is approximately double its size (198kD vs 98kD). Transferrin receptor has two disulfide bonds and exists as a dimer in vivo, so I would expect the undenatured sample to run as a dimer. But what I really want to know is why does it dimerize when heated with BME in SDS sample buffer? A colleague thinks that it just gets too denatured and runs as an aggregate - this person prepares the samples by treating with 2% BME at room temperature for 10 minutes. I have found that if I treat with 5% BME at room temperature I begin to see some of the larger form, but if I boil the sample and leave out the BME I don't see any of the larger form. I suspect that some of the protein modifications are being crosslinked by the BME. Transferrin receptor has 3 N-linked glycosylation sites, 1 O-linked glycosylation site, and is post-translationally phosphorylated and fatty acylated. Also, a colleague told me that a protein she works with, a mannose-6-phosphate receptor present in the golgi, has the same problem with running as a dimer if heated with reductant, but runs fine if heated without reductant. Both proteins are transmembrane receptors and I know the mannose-6-phosphate receptor also has N-linked glycosylation sites. Any ideas?