Force measurements show that uL4 and uL24 mechanically stabilize a fragment of 23S rRNA essential for ribosome assembly.

In vitro reconstitution studies have shown that ribosome assembly is highly cooperative and starts with the binding of a few ribosomal (r-) proteins to rRNA. It is unknown how these early binders act. Focusing on the initial stage of the assembly of the large subunit of the Escherichia coli ribosome, we prepared a 79-nucleotide-long region of 23S rRNA encompassing the binding sites of the early binders uL4 and uL24. Force signals were measured in a DNA/RNA dumbbell configuration with a double optical tweezers setup. The rRNA fragment was stretched until unfolded, in the absence or in the presence of the r-proteins (either uL4, uL24, or both). We show that the r-proteins uL4 and uL24 individually stabilize the rRNA fragment, both acting as molecular clamps. Interestingly, this mechanical stabilization is enhanced when both proteins are bound simultaneously. Independently, we observe a cooperative binding of uL4 and uL24 to the rRNA fragment. These two aspects of r-proteins binding both contribute to the efficient stabilization of the 3D structure of the rRNA fragment under investigation. We finally consider implications of our results for large ribosomal subunit assembly.

Overstretching Double-Stranded RNA, Double-Stranded DNA, and RNA-DNA Duplexes.

Using single-molecule force measurements, we compare the overstretching transition of the four types of duplexes composed of DNA or RNA strands. Three of the four extremities of each double helix are attached to two microscopic beads, and a stretching force is applied with a dual-beam optical trapping interferometer. We find that overstretching occurs for all four duplexes with small differences between the plateau forces. Double-stranded RNA (dsRNA) exhibits a smooth transition in contrast to the other three duplexes that show sawtooth patterns, the latter being a characteristic signature of peeling. This difference is observed for a wide range of experimental conditions. We present a theoretical description that explains the difference and predicts that peeling and bubble formation do not occur in overstretching double-stranded RNA. Formation of S-RNA is proposed, an overstretching mechanism that contrary to the other two does not generate single strands. We suggest that this singular RNA property helps RNA structures to assemble and play their essential roles in the biological cell.

Hairpins under tension: RNA versus DNA.

We use optical tweezers to control the folding and unfolding of individual DNA and RNA hairpins by force. Four hairpin molecules are studied in comparison: two DNA and two RNA ones. We observe that the conformational dynamics is slower for the RNA hairpins than for their DNA counterparts. Our results indicate that structures made of RNA are dynamically more stable. This difference might contribute to the fact that DNA and RNA play fundamentally different biological roles in spite of chemical similarity.

Revealing the competition between peeled ssDNA, melting bubbles, and S-DNA during DNA overstretching using fluorescence microscopy.

Mechanical stress plays a key role in many genomic processes, such as DNA replication and transcription. The ability to predict the response of double-stranded (ds) DNA to tension is a cornerstone of understanding DNA mechanics. It is widely appreciated that torsionally relaxed dsDNA exhibits a structural transition at forces of ∼65 pN, known as overstretching, whereby the contour length of the molecule increases by ∼70%. Despite extensive investigation, the structural changes occurring in DNA during overstretching are still generating considerable debate. Three mechanisms have been proposed to account for the increase in DNA contour length during overstretching: strand unpeeling, localized base-pair breaking (yielding melting bubbles), and formation of S-DNA (strand unwinding, while base pairing is maintained). Here we show, using a combination of fluorescence microscopy and optical tweezers, that all three structures can exist, uniting the often contradictory dogmas of DNA overstretching. We visualize and distinguish strand unpeeling and melting-bubble formation using an appropriate combination of fluorescently labeled proteins, whereas remaining B-form DNA is accounted for by using specific fluorescent molecular markers. Regions of S-DNA are associated with domains where fluorescent probes do not bind. We demonstrate that the balance between the three structures of overstretched DNA is governed by both DNA topology and local DNA stability. These findings enhance our knowledge of DNA mechanics and stability, which are of fundamental importance to understanding how proteins modify the physical state of DNA.

Probing ribosomal protein-RNA interactions with an external force.

Ribosomal (r-) RNA adopts a well-defined structure within the ribosome, but the role of r-proteins in stabilizing this structure is poorly understood. To address this issue, we use optical tweezers to unfold RNA fragments in the presence or absence of r-proteins. Here, we focus on Escherichia coli r-protein L20, whose globular C-terminal domain (L20C) recognizes an irregular stem in domain II of 23S rRNA. L20C also binds its own mRNA and represses its translation; binding occurs at two different sites--i.e., a pseudoknot and an irregular stem. We find that L20C makes rRNA and mRNA fragments encompassing its binding sites more resistant to mechanical unfolding. The regions of increased resistance correspond within two base pairs to the binding sites identified by conventional methods. While stabilizing specific RNA structures, L20C does not accelerate their formation from alternate conformations--i.e., it acts as a clamp but not as a chaperone. In the ribosome, L20C contacts only one side of its target stem but interacts with both strands, explaining its clamping effect. Other r-proteins bind rRNA similarly, suggesting that several rRNA structures are stabilized by "one-side" clamping.

RNA Folding and Unfolding Under Force: Single-Molecule Experiments and Their Analysis.

We have previously described (Geffroy et al. Methods Mol Biol 1665:25-40, 2018) how to unfold (or fold) a single RNA molecule under force using a dual-beam optical trap setup. In this chapter, we complementarily describe how to analyze the corresponding data and how to interpret it in terms of RNA three-dimensional structure. As with all single-molecule methods, single RNA molecule force data often exhibit several discrete states where state-to-state transitions are blurred in a noisy signal. In order to cope with this limitation, we have implemented a novel strategy to analyze the data, which uses a hidden Markov modeling procedure. A representative example of such an analysis is presented.

DNA Hybridization Measured with Graphene Transistor Arrays.

Arrays of field-effect transistors are fabricated from chemical vapor deposition grown graphene (GFETs) and label-free detection of DNA hybridization performed down to femtomolar concentrations. A process is developed for large-area graphene sheets, which includes a thin Al2 O3 layer, protecting the graphene from contamination during photolithographic patterning and a SiOx capping for biocompatibility. It enables fabrication of high-quality transistor arrays, exhibiting stable close-to-zero Dirac point voltages under ambient conditions. Passivation of the as-fabricated chip with a layer composed of two different oxides avoids direct electrochemical contact between the DNA solutions and the graphene layer during hybridization detection. DNA probe molecules are electrostatically immobilized via poly-l-lysine coating of the chip surface. Adsorption of this positively charged polymer induces a positive shift of the Dirac point and subsequent immobilization of negatively charged DNA probes induces a negative shift. Spatially resolved hybridization of DNA sequences is performed on the GFET arrays. End-point as well as real-time in situ measurements of hybridization are achieved. A detection limit of 10 fm is observed for hybridization of 20-nucleotide DNA targets. Typical voltage signals are around 100 mV and spurious drifts below 1 mV per hour.

Probing DNA base pairing energy profiles using a nanopore.

We experimentally show that the voltage driven unzipping of long DNA duplexes by an alpha-hemolysin pore is sensitive to the shape of the base pairing energy landscape. Two sequences of equal global stability were investigated. The sequence with an homogeneous base pairing profile translocates faster than the one with alternative weak and strong regions. We could qualitatively account for theses observations by theoretically describing the voltage driven translocation as a biased random walk of the unzipping fork in the sequence dependent energy landscape.

Single-Molecule FRET Assay to Observe the Activity of Proteins Involved in RNA/RNA Annealing.

In recent years, single-molecule fluorescence resonance energy transfer (smFRET) has emerged as a powerful technique to study macromolecular interactions. The chief advantages of smFRET analysis compared to bulk measurements include the possibility to detect sample heterogeneities within a large population of molecules and the facility to measure kinetics without needing the synchronization of intermediate states. As such, the methodology is particularly well adapted to observe and analyze RNA/RNA and RNA/protein interactions involved in small noncoding RNA-mediated gene regulation networks. In this chapter, we describe and discuss protocols that can be used to measure the dynamics of these interactions, with a particular emphasis on the advantages-and experimental pitfalls-of using the smFRET methodology to study sRNA-based biological systems.

RNA Unzipping and Force Measurements with a Dual Optical Trap.

In order to mechanically unfold a single RNA molecule, an RNA/DNA hybrid construction is prepared which allows specific attachment to two micrometer-sized beads. A dual-beam optical trap thus holding the construct in solution captures the beads separately. Unfolding of a molecule is obtained by increasing the distance between the traps, one trap being slowly moved while the other is held fixed. Force is measured to sub-piconewton precision by back focal plane interferometry of the bead in the fixed trap. The experiment allows us to measure structure and base-sequence-dependent force signals. In this chapter, important technical aspects of this type of single-molecule force measurements are considered.

Single-molecule manipulation of nucleic acids.

During the past decade, local force measurement techniques, such as atomic force microscopy and optical tweezers, were used to study the elastic properties and mechanically induced structural transitions of nucleic acids at the single-molecule level. Single-molecule manipulation has also increasingly been used to investigate DNA-dependent enzymatic processes, with implications for unfolding and modifying DNA, protein-DNA interactions, replication and transcription. Compared to classical techniques of molecular biology, single-molecule measurements avoid the need to average over a large number of events, and can thus potentially provide detailed and complementary information.

Electronic hybridization detection in microarray format and DNA genotyping.

We describe an approach to substituting a fluorescence microarray with a surface made of an arrangement of electrolyte-gated field effect transistors. This was achieved using a dedicated blocking of non-specific interactions and comparing threshold voltage shifts of transistors exhibiting probe molecules of different base sequence. We apply the approach to detection of the 35delG mutation, which is related to non-syndromic deafness and is one of the most frequent mutations in humans. The process involves barcode sequences that are generated by Tas-PCR, a newly developed replication reaction using polymerase blocking. The barcodes are recognized by hybridization to surface attached probes and are directly detected by the semiconductor device.

DNA unzipping and force measurements with a dual optical trap.

In order to open the DNA double helix mechanically, a molecular construction is prepared which allows specific attachment of the two complementary strands of an individual molecule to two different µm-sized beads. The beads are separately captured by a dual optical trap, thus holding the DNA construction in solution. The opening of a molecule is obtained by increasing the distance between the traps, one trap being slowly moved while the other is held fixed. Force is measured to sub-piconewton precision by back focal plane interferometry of the bead in the fixed trap. The experiment allows us to measure base sequence-dependent force signal. In this chapter, important technical aspects of this type of single-molecule force measurements are considered.

Rectification of the current in alpha-hemolysin pore depends on the cation type: the alkali series probed by MD simulations and experiments.

A striking feature of the alpha-hemolysin channel-a prime candidate for biotechnological applications-is the dependence of its ionic conductance on the magnitude and direction of the applied bias. Through a combination of lipid bilayer single-channel recording and molecular dynamics (MD) simulations, we characterized the current-voltage relationship of alpha-hemolysin for all alkali chloride salts at neutral pH. The rectification of the ionic current was found to depend on the type of cations and increase from Li(+) to Cs(+). Analysis of the MD trajectories yielded a simple quantitative model that related the ionic current to the electrostatic potential, the concentration and effective mobility of ions in the channel. MD simulations reveal that the major contribution to the current asymmetry and rectification properties originates from the cationic contribution to the current that is significantly reduced in a cationic dependent way when the membrane polarity is reversed. The variation of chloride current was found to be less important. We report that the differential affinity of cations for the charged residues positioned at the channel's end modulates the number of ions inside the channel stem thus affecting the current properties. Through direct comparison of simulation and experiment, this study evaluates the accuracy of the MD method for prediction of the asymmetric, voltage dependent conductances of a membrane channel.

Interference and crosstalk in double optical tweezers using a single laser source.

Experimental studies of single molecule mechanics require high force sensitivity and low drift, which can be achieved with optical tweezers. We built an optical tweezer setup for force measurements in a two bead assay. A cw infrared laser beam is split by polarization and focused by a high numerical aperture objective to create two traps. The same laser is used to form both traps and to measure the force by back focal plane interferometry. We show that although the two beams entering the microscope are designed to exhibit orthogonal polarization, interference and a significant parasitic force signal occur. Comparing the experimental results with a ray optics model, we show that the interference patterns are caused by the rotation of polarization on microscope lens surfaces and slides. The model qualitatively describes the pattern and the dependence of the parasitic force signal on the experimental parameters. We present two different approaches to experimentally reduce the crosstalk, namely, polarization rectification and frequency shifting.

Rotational drag on DNA: a single molecule experiment.

Within a single-molecule configuration, we have studied rotational drag on double stranded linear DNA by measuring the force during mechanical opening and closing of the double helix at different rates. The molecule is cranked at one end by the effect of unzipping and is free to rotate at the other end. In this configuration the rotational friction torque tau on double-stranded DNA leads to an additional contribution to the opening force. It is shown that the effect of rotational drag increases with the length of the molecule, is approximately proportional to the angular velocity of cranking, and we estimate that the torque tau is of the order of 1k(B)T for 10 000 base pairs of DNA cranked at 2000 turns per second.