Inside every living organism, a plethora of molecular motors are constantly using chemical energy to power cellular processes and perform mechanical tasks necessary for the proper functioning of the cell. How these motors achieve these functions in highly fluctuating environments and the mechanisms underlying their efficient force generation in energy regimes close to the thermal fluctuations remain active questions in modern biophysics. A detailed understanding of the operation of these motors requires access to their molecular trajectories (i.e.: their step-wise motion as a function of time) as well as precise measurement of the forces at which they operate. Single-molecule manipulation methods such as optical tweezers can provide the spatio-temporal resolution necessary to study the translocation of most of these motors whose step sizes are typically in the range of a few nanometers. However, for some molecular motors, particularly those with the shortest step-size reported so far, a single DNA or RNA basepair (~3.4 Å), their molecular trajectories remain difficult to obtain. Therefore, finding experimental conditions that improve the spatio-temporal resolution, robustness, and accuracy of optical tweezers can pave the way to a better understanding of the mechanochemistry of such motors.

RNA polymerases (RNAP), the central enzymes of transcription thus gene expression, are responsible for converting the stored information in DNA into active form as an RNA transcript. During transcription elongation, RNA polymerases undergo multiple nucleotide addition cycles in which a cognate nucleotide triphosphate (NTP) binds to the active site of the enzyme, the NTP is subsequently incorporated into RNA by catalysis, and finally, RNAP translocates by one base pair (~3.4 Å) along the DNA to restart the cycle. The robust detection of this tiny movement of RNA polymerases moving along the DNA and the measurement of the dwell time of individual nucleotide cycles have remained elusive to biophysicists. In this dissertation, I describe our efforts to improve the spatio-temporal resolution of optical tweezers to reliably detect these small displacements. A systematic characterization of the noise in two high resolution dual-trap optical tweezers setups, differential-path and time-shared, is described. The improved spatio-temporal resolution of time-shared dual-trap optical tweezers in combination with improved step-finding algorithms allowed us to robustly observe the molecular trajectories of E. coli RNA polymerase at single base-pair resolution over long distances, and thus record its distribution of the dwell times.

Despite the unique information about the trajectories of molecular motors that can be obtained using optical tweezers, the inherent one-dimensional nature of these measurements only allow access to finite information such as the total dwell time of a motor with further details of their mechanochemistry usually only obtained by indirect experiments. Therefore, a combination of optical tweezers with an orthogonal readout such as single-molecule fluorescence can lead to new insights on the mechanisms of molecular motors which could be hardly obtained if these experiments were done in isolation. We developed experimental conditions to monitor the active translocation of dual fluorescently-labeled RNA polymerase using an optical tweezers instrument with fluorescence capabilities (“Fleezers”). By monitoring the conformational changes of the trigger-loop of E. coli RNA polymerase by single-molecule FRET (smFRET) and the translocation of the enzyme by optical tweezers, we were able to directly detect the precise moment of nucleotide triphosphate binding, catalysis, and translocation of RNA polymerase in real-time at the single-molecule level.

We further exploited the capabilities of the Fleezers setup for the study of other biological systems important to the process of transcription. Nucleosomes, the fundamental structural unit of eukaryotic chromatin, are constantly subject to passive spontaneous fluctuations and active forces generated by polymerases and chromatin remodelers that lead to nucleosomal DNA unwrapping/rewrapping. Despite numerous studies on this process using single-molecule manipulation methods, questions regarding the actual conformational changes happening during nucleosomal DNA unwrapping, as well as the relationship between DNA unwrapping and the integrity of the histone protein core, remained unanswered. By simultaneously monitoring the integrity of nucleosomes using fluorescently labeled H2A nucleosomes, we unveiled a causal relationship between nucleosomal DNA unwrapping and the loss of H2A/H2B dimers during their mechanical manipulation. Similarly, a simultaneous measurement of the dynamics of nuclesomal DNA arms by smFRET and their corresponding force/extension transitions allowed us to describe the asymmetric conformational changes that nucleosomes undergo during their unwrapping/rewrapping.

Finally, I briefly describe in this dissertation, preliminary results concerning the development of methodologies to study the role of topology on DNA dynamics and transcription. DNA inside living organisms is topologically constrained leading to considerable deviations from its canonical B-form that ultimately affect the way in which DNA-binding proteins interact with it. Despite revealing important features of the transcription process, in vitro studies are usually performed on non-topologically constrained linear DNA molecules. Thus, it is not clear how the observed transcriptional process could change if DNA topology were to be introduced. Some of the current limitations to perform biophysical studies on DNA topology concern the challenges to generate suitable DNA samples with specific label positions and to manipulate their topology in a controllable way. In this dissertation, a method to site-specifically label circular DNA molecules and to control their topology by enzymatic treatment is introduced. This novel approach allowed us to perform biophysical measurements to study the role of DNA topology on transcription.