Abstract
Self-assembled monolayers (SAMs) of alkanethiolates on gold have become an important tool for probing cell-material interactions. Emerging studies in stem cell biology are particularly reliant on well-defined model substrates, and rapid, highly controllable fabrication methods may be necessary for characterizing the wide array of stem cell-material interactions. Therefore, this study describes a rapid method for creating SAM cell culture substrates with multiple discrete regions of controlled peptide identity and density. The approach uses a NaBH(4) solution to selectively remove regions of bioinert, hydroxyl-terminated oligo(ethylene glycol) alkanethiolate SAM and then locally replace them with mixed SAMs of hydroxyl- and carboxylic acid-terminated oligo(ethylene glycol) alkanethiolates. The cell adhesion peptide Arg-Gly-Asp-Ser-Pro (RGDSP) was then covalently linked to carboxylic acid-terminated mixed SAM regions to create cell adhesive environments within a bioinert background. SAM preparation and peptide immobilization were characterized using polarization modulation-infrared reflection-absorption spectroscopy (PM-IRRAS), as well as assays to monitor conjugation of a fluorescently labeled peptide. This "localized SAM replacement" method was achieved using an array of microchannels, which facilitated rapid and simple processing. Results indicate that immobilized RGDSP promoted spatially localized attachment of human mesenchymal stem cells (hMSCs) within specified regions, while maintaining a stable, bioinert background in serum-containing cell culture conditions for up to 14 days. Cell attachment to patterned regions presenting a range of cell adhesion peptide densities demonstrated that peptide identity and density strongly influence hMSC spreading and focal adhesion density. These substrates contain discrete, well-defined microenvironments for stem cell culture, which could ultimately enable high-throughput screening for the effects of immobilized signals on stem cell phenotype.