I have been having issues with non-specific protein adhesion to my PEG-monolayer that I have formed.
First things first, The surface was first cleaned using Nanostrip (stabilized piranha), and sonicated in DI for 1hr. The surface was then dried under a nitrogen stream. I then used a 2% solution of PEG-Silane (MW ~700from Gelest Inc.) in toluene, and allowed to react under nitrogen for 16 hours.
I find that the monolayer works well at resisting some protein adhesion. For example, I can prevent streptavidin from adhering to the surface. But sticker proteins, such as laminin or PDL (Poly-lysine) are non-specifically adhering to the background. I have seen very little literature on this matter of different proteins behaving differently on PEG-monolayers.
I also tried a glutaraldehyde crosslinking method of PEG-NH2 to an APTES monolayer, and a nucleophilic substitution method using PEG-NHS onto an APTES monolayer. None of these worked to prevent the adhesion either.
I cannot use a BSA blocking method, as my aim is to avoid cell adhesion, and my cell cultures are sticking to BSA passivated surfaces.
Has anyone experienced this before?
mascott wrote:
I'm not working with cells, but I've done something similar. I used APTMS(basically the same as APTES) and then I used PEG5-4FB-NHS(from Solulink, used at 350 uM in 150 mM NaCL, 100 mM PBS, pH 7.2 overnight) on the surface, and then 0.02% tween-20 in the running buffer. Under these conditions, my polyclonal IgG-HRP @ 0.2 mg/mL didn't stick. I don't know if that amount of tween is compatible with your cells, but you could probably go still lower. Another thing you should try is the silanization at 70 C for one hour, with continuous gentle agitation. I've gotten the most uniform monolayers that way, so at least that removes the roughness that cells like to stick to, and I imagine with a 16 hour reaction you're getting a fairly rough multilayer surface, and not a monolayer at all.
See Pasternack et al. for more info on this method.