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Single-molecule tracking (SMT) is a powerful tool for real-time studies of protein interactions in living cells. Dye-labelled SNAP-tag and HaloTag self-labelling proteins have simplified SMT significantly, due to their superior photophysical properties compared to fluorescent proteins. However, due to their size, fusion of these tags to a protein of interest often results in loss of protein function. We introduce FLORENCE – a universal labelling method for SMT, based on genetic code expansion (GCE). We overcome significant caveats related to re-coded strains, vectors, and dyes and report successful tracking of site-specifically intracellularly labelled proteins in genomically re-coded E. coli. Our findings establish a robust in vivo protein-labelling strategy, expanding the capabilities of SMT as a method to study the dynamics of proteins in living cells. Moreover, we observe that the strain-promoted azide–alkyne click-chemistry reaction occurs as fast as 30 min in live E. coli cells and can be used as a robust labelling reaction.

Electroporation of dye-labeled bio-molecules into bacteria has proven to be a valuable route for single-molecule tracking in living cells. However, control over cell viability, electroporation efficiency, and environment conditions before, during, and after electroporation is difficult to achieve in bulk experiments. Here, a microfluidic platform is presented capable of single-cell electroporation with in situ microscopy and demonstrate delivery of DNA into bacteria. Via real time observation of the electroporation process, it is found that the effect of electrophoresis plays an important role when performing electroporation in a miniaturized platform and show that its undesired action can be balanced by using bipolar electrical pulses. It is suggested that a low temperature of the sample during electroporation is important for cell viability due to temperature-dependant viscoelastic properties of the cell membrane. It is further found that the presence of low conductive liquid between cells and the electrodes leads to a voltage divider effect that strongly influences the success of on-chip electroporation. Finally, it is concluded that electroporation is a highly stochastic process and envision that the microfluidic system presented here, capable of single-cell read-out, can be used for further fundamental studies to increase the understanding of the electroporation process in bacterial cells.

The bacterial chaperone Trigger factor (TF) binds to ribosome-nascent chain complexes (RNCs) and cotranslationally aids the folding of proteins in bacteria. Decades of studies have given a broad, but often conflicting, description of the substrate specificity of TF, its RNC-binding dynamics, and competition with other RNC-binding factors, such as the Signal Recognition Particle (SRP). Previous RNC-binding kinetics experiments were commonly conducted on stalled RNCs in reconstituted systems, and consequently, may not be representative of the interaction of TF with ribosomes translating mRNA in the cytoplasm of the cell. Here, we used single-particle tracking (SPT) to measure TF binding to actively translating ribosomes inside living Escherichia coli. In cells, TF displays distinct binding modes—longer (ca 1 s) and shorter (ca 50 ms) RNC bindings. Consequently, we conclude that TF, on average, stays bound to the RNC for only a fraction of the translation cycle. Further, binding events are interrupted only by transient excursions to a freely diffusing state (ca 40 ms), suggesting a highly dynamic binding and unbinding cycle of TF in vivo. We also show that TF competes with SRP for RNC binding, and in doing so, tunes the binding selectivity of SRP.

Metelev et al. use single-molecule tracking to study kinetics of translation directly in E. coli cells, and how it is affected by translation inhibitors and rRNA mutations. Their results support widespread 70S re-initiation on mRNAs. Ribosome mediated mRNA translation is central to life. The cycle of translation, however, has been characterized mostly using reconstituted systems, with only few techniques applicable for studies in the living cell. Here we describe a live-cell ribosome-labeling method, which allows us to characterize the whole processes of finding and translating an mRNA, using single-molecule tracking techniques. We find that more than 90% of both bacterial ribosomal subunits are engaged in translation at any particular time, and that the 30S and 50S ribosomal subunits spend the same average time bound to an mRNA, revealing that 30S re-initiation on poly-cistronic mRNAs is not prevalent in E. coli. Instead, our results are best explained by substantial 70S re-initiation of translation of poly-cistronic mRNAs, which is further corroborated by experiments with translation initiation inhibitors. Finally, we find that a variety of previously described orthogonal ribosomes, with altered anti-Shine-Dalgarno sequences, show significant binding to endogenous mRNAs.

Mechanistic details of the signal recognition particle (SRP)-mediated insertion of membrane proteins have been described from decades of in vitro biochemical studies. However, the dynamics of the pathway inside the living cell remain obscure. By combining in vivo single-molecule tracking with numerical modeling and simulated microscopy, we have constructed a quantitative reaction-diffusion model of the SRP cycle. Our results suggest that the SRP-ribosome complex finds its target, the membrane-bound translocon, through a combination of three-dimensional (3D) and 2D diffusional search, together taking on average 750 ms. During this time, the nascent peptide is expected to be elongated only 12 or 13 amino acids, which explains why, in Escherichia coli, no translation arrest is needed to prevent incorrect folding of the polypeptide in the cytosol. We also found that a remarkably high proportion (75%) of SRP bindings to ribosomes occur in the cytosol, suggesting that the majority of target ribosomes bind SRP before reaching the membrane. In combination with the average SRP cycling time, 2.2 s, this result further shows that the SRP pathway is capable of targeting all substrate ribosomes to translocons.

The spread of antibiotic resistance is turning many of the currently used antibiotics less effective against common infections. To address this public health challenge, it is critical to enhance our understanding of the mechanisms of action of these compounds. Aminoglycoside drugs bind the bacterial ribosome, and decades of results from in vitro biochemical and structural approaches suggest that these drugs disrupt protein synthesis by inhibiting the ribosome's translocation on the messenger RNA, as well as by inducing miscoding errors. So far, however, we have sparse information about the dynamic effects of these compounds on protein synthesis inside the cell. In the present study, we measured the effect of the aminoglycosides apramycin, gentamicin, and paromomycin on ongoing protein synthesis directly in live Escherichia coli cells by tracking the binding of dye-labeled transfer RNAs to ribosomes. Our results suggest that the drugs slow down translation elongation two- to fourfold in general, and the number of elongation cycles per initiation event seems to decrease to the same extent. Hence, our results imply that none of the drugs used in this study cause severe inhibition of translocation.

Decades of traditional biochemistry, structural approaches, and, more recently, single-molecule-based in vitro techniques have provided us with an astonishingly detailed understanding of the molecular mechanism of ribosome-catalyzed protein synthesis. However, in order to understand these details in the context of cell physiology and population biology, new techniques to probe the dynamics of molecular processes inside the cell are needed. Recent years' development in super-resolved fluorescence microscopy has revolutionized imaging of intracellular processes, and we now have the possibility to directly peek into the microcosm of biomolecules in their native environment. In this Perspective, we discuss how these methods are currently being applied and further developed to study the kinetics of protein synthesis directly inside living cells.

Chloramphenicol is a broad-spectrum antibiotic targeting the protein synthesis machinery by binding to the bacterial ribosome. Chloramphenicol has been considered a classic general inhibitor of translation, blocking the accommodation of aa-tRNA into the A site of the large ribosomal subunit. However, recent studies suggest that this proposed mechanism is a simplification and that the effect of chloramphenicol on mRNA translation is much more dynamic. By tracking single dye-labelled elongator and initiator tRNAs in Escherichia coli cells treated with chloramphenicol, we observe the direct effect of chloramphenicol on translation kinetics. We find clear indications of slow but significant mRNA translation on drug bound ribosomes.

Our ability to directly relate results from test-tube biochemical experiments to the kinetics in living cells is very limited. Here we present experimental and analytical tools to directly study the kinetics of fast biochemical reactions in live cells. Dye-labeled molecules are electroporated into bacterial cells and tracked using super-resolved single-molecule microscopy.Trajectories are analyzed by machine-learning algorithms to directly monitor transitions between bound and free states. In particular, we measure the dwell time of tRNAs on ribosomes, and hence achieve direct measurements of translation rates inside living cells at codon resolution. We find elongation rates with tRNA(Phe) that are in perfect agreement with previous indirect estimates, and once fMet-tRNA(fMet) has bound to the 30S ribosomal subunit, initiation of translation is surprisingly fast and does not limit the overall rate of protein synthesis. The experimental and analytical tools for direct kinetics measurements in live cells have applications far beyond bacterial protein synthesis.