Fluorescent Labeling of Proteins in Whole Cell Extracts for Single-Molecule Imaging.

Cellular machines such as the spliceosome and ribosome can be composed of dozens of individual proteins and nucleic acids. Given this complexity, it is not surprising that many cellular activities have not yet been biochemically reconstituted. Such processes are often studied in vitro in whole cell or fractionated lysates. This presents a challenge for obtaining detailed biochemical information when the components being investigated may be only a minor component of the extract and unrelated processes may interfere with the assay. Single-molecule fluorescence microscopy methods allow particular biomolecules to be analyzed even in the complex milieu of a cell extract. This is due to the use of bright fluorophores that emit light at wavelengths at which few cellular components fluoresce, and the development of chemical biology tools for attaching these fluorophores to specific cellular proteins. Here, we describe a protocol for fluorescent labeling of endogenous, SNAP-tagged yeast proteins in whole cell extract. This method allows biochemical reactions to be followed in cell lysates in real time using colocalization single-molecule fluorescence microscopy. Labeled complexes can also be isolated from extract and characterized by SNAP tag single-molecule pull-down (SNAP-SiMPull). These approaches have proven useful for studying complex biological machines such as the spliceosome that cannot yet be reconstituted from purified components.

Evolution and characterization of a benzylguanine-binding RNA aptamer.

Repurposing the "protein-labeling toolkit" for RNA research could be a pragmatic approach for developing new RNA-labeling methods. We have evolved an RNA aptamer that tightly binds benzylguanine (bG), the key ligand for the protein SNAP-tag. The aptamer tightly binds bG fluorophores and can be purified from cellular RNA with bG agarose under native conditions.