Förster resonance energy transfer (FRET) spectrometry is a method for determining the quaternary structure of protein oligomers from distributions of FRET efficiencies that are drawn from pixels of fluorescence images of cells expressing the proteins of interest. FRET spectrometry protocols currently rely on obtaining spectrally resolved fluorescence data from intensity-based experiments. Another imaging method, fluorescence lifetime imaging microscopy (FLIM), is a widely used alternative to compute FRET efficiencies for each pixel in an image from the reduction of the fluorescence lifetime of the donors caused by FRET. In FLIM studies of oligomers with different proportions of donors and acceptors, the donor lifetimes may be obtained by fitting the temporally resolved fluorescence decay data with a predetermined number of exponential decay curves. However, this requires knowledge of the number and the relative arrangement of the fluorescent proteins in the sample, which is precisely the goal of FRET spectrometry, thus creating a conundrum that has prevented users of FLIM instruments from performing FRET spectrometry. Here, we describe an attempt to implement FRET spectrometry on temporally resolved fluorescence microscopes by using an integration-based method of computing the FRET efficiency from fluorescence decay curves. This method, which we dubbed time-integrated FRET (or tiFRET), was tested on oligomeric fluorescent protein constructs expressed in the cytoplasm of living cells. The present results show that tiFRET is a promising way of implementing FRET spectrometry and suggest potential instrument adjustments for increasing accuracy and resolution in this kind of study.

Many biomolecular condensates, including nucleoli and stress granules, form via dynamic multivalent protein-protein and protein-RNA interactions. These molecular interactions nucleate liquid-liquid phase separation (LLPS) and determine condensate properties, such as size and fluidity. Here, we outline the experimental procedures for single-molecule fluorescence experiments to probe protein-RNA interactions underlying LLPS. The experiments include single-molecule Förster (Fluorescence) resonance energy transfer (smFRET) to monitor protein-induced conformational changes in the RNA, protein-induced fluorescence enhancement (PIFE) to measure protein-RNA encounters, and single-molecule nucleation experiments to quantify the association and buildup of proteins on a nucleating RNA. Together, these experiments provide complementary approaches to elucidate a molecular view of the protein-RNA interactions that drive ribonucleoprotein condensate formation.

Single molecule fluorescence and nuclear magnetic resonance spectroscopy (NMR) are two very powerful techniques for the analysis of intrinsically disordered proteins (IDPs). Both techniques have individually made major contributions to deciphering the complex properties of IDPs and their interactions, and it has become evident that they can provide very complementary views on the distance-dynamics relationships of IDP systems. We now review the first approaches using both NMR and single molecule fluorescence to decipher the molecular properties of IDPs and their interactions. We shed light on how these two techniques were employed synergistically for multidomain proteins harboring intrinsically disordered linkers, for veritable IDPs, but also for liquid-liquid phase separated systems. Additionally, we provide insights into the first approaches to use single molecule Förster resonance energy transfer (FRET) and NMR for the description of multiconformational models of IDPs.

To identify the translocation components in cells, and to understand how they function in protein transport and membrane insertion, a variety of techniques have been used such as genetics, biochemistry, structural biology and single molecule methods. In particular, site-directed crosslinking between the client proteins and components of the translocation machineries have contributed significantly in the past and will do so in the future. One advantage of this technology is that it can be applied in vivo as well as in vitro and a comparison of the two approaches can be made. Also, the in vivo techniques allow time-dependent protocols which are essential for studying cellular pathways. Protein purification and reconstitution into proteoliposomes are the gold standard for studying membrane-based transport and translocation systems. With these biochemically defined approaches the function of each component in protein transport can be addressed individually with a plethora of biophysical techniques. Recently, the use of nanodiscs for reconstitution has added another extension of this reductionistic approach. Fluorescence based studies, cryo-microscopy and NMR spectroscopy have significantly added to our understanding how proteins move into and across membranes and will do this also in future.