Interactions between cytosolic biomolecules and the bacterial inner membrane are fundamental to many cellular processes, yet directly measuring their binding kinetics in living cells remains challenging. Conventional 2D single-molecule tracking analyses can be insufficient, particularly when membrane association does not markedly alter the diffusion rate. Here, we present a method to recover membrane interaction kinetics from 3D single-molecule trajectories in rod-shaped bacteria. Using simulated 3D tracking data, we identify membrane-associated motion by quantifying how well short trajectory segments follow the circular curvature of the cell membrane. The resulting measure is further analyzed using a hidden Markov modeling framework, enabling robust discrimination between cytosolic and membrane-bound states and capturing the dynamics of state transitions without requiring diffusion-rate changes or direct colocalization with membrane markers. This work establishes a general framework for extracting membrane interaction kinetics from 3D single-molecule tracking data in live bacteria and highlights the value of realistic microscopy simulations for quantitative interpretation and systematic bias assessment.

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.

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.

With recent years' revolution in super-resolution imaging of sub-cellular physiology and dynamics, we have now reached the point when quantitative biochemical experiments can be performed directly inside living cells rather than in the test tube. In our lab, we are developing tools to study spatiotemporal dynamics of protein synthesis, targeting and folding in live E. coli cells. With single-molecule tracking techniques in combination with hidden-Markov modelling based analysis of diffusion trajectories, we follow labelled molecules through their complete binding cycles, and thereby extract reaction kinetics data.

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.

Electroporation is a biophysical phenomenon in which membrane pores are created under the conditions of an externally applied electrical field. It is used for a wide variety of purposes, ranging from applications in the food industry to scientific research. In microbiology, electroporation is routinely used for gene transfection in bacteria, such as the delivery of plasmid DNA. In practice, this is achieved by applying a voltage of a few kV to the cell suspension in electroporation cuvettes with a 1–2 mm gap between the electrodes.

The bacterial chaperone Trigger factor (TF) binds to ribosome-nascent chain complexes (RNCs) and co-translationally 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 conducted on stalled RNCs in reconstituted systems, and consequently, may not represent 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 two distinct binding modes — long (ca 1 s) target-specific RNC binding, and shorter (ca 50 ms) sampling of non-target RNCs. RNC binding events are interrupted only by transient excursions to a freely diffusing state (ca 40 ms). We also show that TF competes with SRP for RNC binding in vivo , and in doing so, tunes the binding selectivity of SRP. ### Competing Interest Statement The authors have declared no competing interest.

Over the decades, reconstituted systems have been exploited for kinetics measurements of biological processes in vitro. However, since in live cells, different biochemical processes are always inter-connected and taking place in a confined geometry, studies of an isolated system can never account for the whole picture. On the other hand, access to kinetics of dynamic processes directly inside the cell has been problematic since the molecules of interest are normally not synchronized with respect to binding state. This problem can be solved by studying individual molecules, one at a time. In the present study we have developed a single-molecule-tracking system to study the kinetics of Signal Recognition Particle (SRP) -mediated targeting of ribosomes to the inner membrane peptide translocation complexes, the translocons. By exploiting electroporation of labeled nucleic acids in live E. coli cells we delivered dye-labeled 4.5S RNA (the RNA component of SRP) into living bacteria. By using new generation self-healing fluorophore (Cy5 analog coupled to triplet quencher) combined with mild oxygen concentration in the cell medium, we achieved superior photostability of labels. We have tracked single SRP through transitions between their different binding states, which allowed us to measure directly, with high precision, spatiotemporal kinetics of the complete SRP cycle. Experimental tracking results were further validated and refined using numerical modelling and state-of-the-art microscopy simulations.

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 use of single-molecule fluorescence microscopy makes it possible to observe molecules of interest in their native environment, one at a time. In this presentation we show how methods recently developed in our lab are applied to study different aspects of protein synthesis kinetics in live bacterial cells. To study translation initiation and elongation kinetics in E.coli, we label both ribosomal subunits with bright and photostable cell-permeable fluorophores. Analysis of single-molecule trajectories enable us to distinguish freely diffusing subunits from translating ribosomes. The results reveal translation kinetics for wt and mutant ribosomes. To shed light on co-translational insertion of inner membrane proteins, we study the dynamics of membrane-targeting of ribosomes by the Signal Recognition Particle (SRP). We deliver dye labeled 4.5S RNA (part of SRP) to E. coli by electroporation, and follow the dynamics of SRP binding to ribosomes and targeting to the membrane. We are further using our previously developed tRNA tracking approach, to study, in situ, the mechanism of action of ribosome-targeting antibiotics, such as erythromycin and aminoglycosides. The analysis of single-molecule trajectories is performed using a Hidden Markov Model (HMM) approach, distinguishing discrete diffusional states and transition rates between them. We are also continuously evaluating our experimental and analytical approach by producing simulated microscopy data, based on reaction-diffusion models in a cell geometry, and running these data through our analysis pipeline.

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.

ABSTRACT Ribosome mediated mRNA translation is central to life as we know it. The cycle of translation has, however, not been characterized in a living cell. Here we have developed a live-cell ribosome-labeling method, which allows us to characterize the whole processes of finding an mRNA and translating it, using single-molecule tracking techniques. We find that more than 90% of both bacterial ribosomal subunits are engaged in elongation at any particular time, and that neither of the subunits, in general, continues translation from one open reading frame to the next on a poly-cistronic mRNA. Furthermore, we find that a variety of previously published orthogonal ribosomes, with altered anti-Shine-Dalgarno sequences, show significant binding to endogenous mRNAs, with the rate of translation initiation only modestly affected. Hence, our results suggest that other mRNA elements than the SD sequence play major roles in directing the ribosome to the correct translation start sites.

Though ribosome catalyzed protein synthesis has been studied extensively over the decades, some blind spots still exist, in particular with respect to reaction kinetics. When it comes to in vivo measurements, where molecules are not synchronized, new single-molecule methods are needed. In our pilot study we applied a single-molecule tracking approach to follow dye-labeled initiator and elongator tRNAs, introduced in E. coli cells by electroporation. Trajectories detected with high spatial and temporal resolution were analysed with a Hidden Markov Model approach allowing extraction of information about the diffusional behaviour of the molecules under investigation. We were able to monitor the dwell times of the tRNAs in different diffusional states, hence directly measure translation initiation and elongation rates1. The elongation rate with tRNAPhe was found to be in perfect agreement with previous indirect estimates. Initiation of translation, once fMet-tRNAfMet has bound to the 30S ribosomal subunit, turned out to be surprisingly fast and does probably not limit the overall protein synthesis rate. We are now applying our developed experimental and analytical tool to other studies. For example, we are investigating the kinetics of Signal Recognition Particle (SRP) mediated co-translational targeting of membrane proteins. 1 Volkov et al, 2018, Nat Chem Biol