I've been trying to pull down a GFP-tagged protein using the Pierce Crosslink Immunoprecipitation kit. Coomassie staining does not seem to have been sensitive enough to detect protein in the eluent so I've been using western blot with the same anti-GFP polyclonal antibody to detect protein bands. In the lysate extract which did not bind to the columns, I am detecting a band corresponding to my protein. I am then eluting using the kit-provided elution buffer, followed by a separate, much harsher elution by boiling the column with SDS/beta-mercaptoethanol gel loading buffer. However, whilst I see a very faint band in the SDS-eluent corresponding to the protein I want (~80kDa), I see a much stronger band at ~50kDa, which is entirely absent in the non-binding lysate sample. Furthermore, using the kit eluent, I only see this ~50kDa band (although this may just be due to the ~80kDa band being too faint to see).
Could anyone explain to me where 30kDa of my protein has gone? A protease inhibitor cocktail was used and steps were carried out at 4 degrees or on ice, except for final elution which only took around 5-10mins. Furthermore, the cell lysate (pre-cleared with control beads) was incubated with the antibody beads for 24 hours, whereas the elution only took well under an hour, so it's strange that such a change would occur in the small amount of time after the two samples were separated. Protease inhibitors were not added to final washing and elution steps, but these were very brief and presumably even if any proteases were present, they would have largely been washed out during the washing stages.
Any help much appreciated!