The SARS-CoV-2 nucleocapsid protein (Np) is essential for viral RNA replication and genomic RNA packaging. Phosphorylation of Np within its central Ser-Arg-rich (SRR) linker is proposed to modulate these functions. To gain mechanistic insights into these distinct roles, we performed in vitro biophysical and biochemical studies using recombinantly expressed ancestral Np and phosphomimetic SRR variants. Limited-proteolysis showed minor cleavage differences between wild-type (WT) and phosphomimetic Np, but no major structure or stability changes in the N- and C-terminal domains were observed by circular dichroism spectroscopy and differential scanning fluorimetry, respectively. Mass photometry (MP) revealed that WT Np dimerized more readily than phosphomimetic variants. Crosslinking-MP showed that WT Np formed discrete complexes on viral 5' UTR stem-loop (SL) 5 RNA, whereas phosphomimetic Np assembled preferentially on SL1-4. WT Np bound non-specifically to all RNAs tested primarily via hydrophobic interactions, whereas phosphomimetic Np showed selectivity for SARS-CoV-2-derived RNAs despite binding more electrostatically. A major difference was observed in the binding kinetics; WT Np compacted and irreversibly bound single-stranded DNA, whereas phosphomimetic Np displayed reduced compaction and fast on/off binding kinetics. These mechanistic insights support a model where phosphorylated Np functions in RNA replication and chaperoning, while non-phosphorylated Np facilitates genomic RNA packaging. The findings also help to explain infectivity differences and clinical outcomes associated with SRR linker variants.

RNA and DNA hairpin formation and disruption play key regulatory roles in a variety of cellular processes. The 59-nucleotide transactivation response (TAR) RNA hairpin facilitates the production of full-length transcripts of the HIV-1 genome. Yet the stability of this long, irregular hairpin becomes a liability during reverse transcription as 24 base pairs must be disrupted for strand transfer. Retroviral nucleocapsid (NC) proteins serve as nucleic acid chaperones that have been shown to both destabilize the TAR hairpin and facilitate strand annealing with its complementary DNA sequence. Yet it has remained difficult to elucidate the way NC targets and dramatically destabilizes this hairpin while only weakly affecting the annealed product. In this work, we used optical tweezers to measure the stability of TAR and found that adding NC destabilized the hairpin and simultaneously caused a distinct change in both the height and location of the energy barrier. This data was matched to an energy landscape predicted from a simple theory of definite base pair destabilization. Comparisons revealed the specific binding sites found by NC along the irregular TAR hairpin. Furthermore, specific binding explained both the unusual shift in the transition state and the much weaker effect on the annealed product. These experiments illustrate a general method of energy landscape transformation that exposes important physical insights.

The L1 retrotransposon is the dominant transposable element in mammalian genomes. L1 comprises at least 20% of the human genome. While most L1 regions are inactive, a few still retain the ability to retrotranspose. L1 encodes two proteins, ORF1p and ORF2p, which are required for retrotransposition. During retrotransposition, ORF2p functions as the reverse transcriptase and the endonuclease. ORF1p is a nucleic acid chaperone that binds nucleic acids with high affinity. However, to date, a detailed mechanistic understanding of ORF1p function in L1 retrotransposition is lacking. The single molecule DNA stretching methods described here have been extensively used to understand ORF1p’s complex nucleic acid binding properties. By correlating these properties to ORF1p’s ability to support L1 retrotransposition in in vivo cell-culture based assays, these studies have significantly contributed to advance the understanding of ORF1p function. Although described in the context of ORF1p, these methods provide a general mechanism to study complex protein-DNA interactions.

Sliding clamp proteins play central roles in DNA metabolism, including replication and repair. The ring-shaped E. coli beta clamp accommodates double-stranded DNA and serves as a platform for proteins involved in multiple DNA transactions. The inner pore of the beta clamp harbors a series of positively charged and polar residues that can bind to the negatively charged backbone of the DNA. These residues are arrayed so that they do not align with the charged phosphates of the DNA backbone. It is hypothesized that this arrangement of these residues provides for the movement of the clamp on DNA as it alternates which residues are bound to the DNA backbone. In this work, we mutated specific charged and polar residues that project into the inner pore of the beta clamp. The beta clamp variants are dimers and have similar thermal stability and in general a similar ability to complement a temperature sensitive strain for growth. One exception was beta-Q149A, which appeared as higher-order species on a native gel although its hydrogen-deuterium exchange pattern measured by mass spectrometry was overall similar to WT beta. These variants all had decreased binding to DNA after loading. Optical tweezers experiments were used to monitor loading on single DNA molecules and measure the rate of beta clamp sliding on DNA. Consistent with the hypothesized role of positively charged residues in the beta inner pore, mutation of one residue resulted in a faster rate of sliding on DNA.

Octahedral rhodium complexes serve as a three-dimensional scaffold for small DNA binding moieties. We compare the DNA binding characteristics of two ligands complexed to rhodium, phenanthrene diimine quinone (phi) and 5,6-chrysene quinone diimine (chrysi). These flat aromatic ligands differ only by an additional ring on chrysi absent in the phi complex. Previous work showed that although chrysi is principally bound to mismatched DNA via metalloinsertion, phi acted as a classical intercalator, binding strongly between basepairs of matched DNA. In dual-beam optical tweezers, we force-unfold DNA hairpins containing key mismatches. We confirm that chrysi stabilizes basepair mismatches, even at zero force, supporting the preferential binding to mismatches through minor groove binding observed in previous studies. However, in contrast to prior work, force extension and constant force data on long, fully paired DNA reveal that both compounds intercalate into double-stranded DNA. Interestingly, chrysi exhibits a higher binding affinity, though the binding kinetics for phi are faster, suggesting tighter binding into the major groove.

Single-stranded nucleic acid (ssNA) binding proteins must both stably protect ssNA transiently exposed during replication and other NA transactions, and also rapidly reorganize and dissociate to allow further NA processing. How these seemingly opposing functions can coexist has been recently elucidated by optical tweezers (OT) experiments that isolate and manipulate single long ssNA molecules to measure conformation in real time. The effective length of an ssNA substrate held at fixed tension is altered upon protein binding, enabling quantification of both the structure and kinetics of protein–NA interactions. When proteins exhibit multiple binding states, however, OT measurements may produce difficult to analyze signals including non-monotonic response to free protein concentration and convolution of multiple fundamental rates. In this review we compare single-molecule experiments with three proteins of vastly different structure and origin that exhibit similar ssNA interactions. These results are consistent with a general model in which protein oligomers containing multiple binding interfaces switch conformations to adjust protein:NA stoichiometry. These characteristics allow a finite number of proteins to protect long ssNA regions by maximizing protein–ssNA contacts while also providing a pathway with reduced energetic barriers to reorganization and eventual protein displacement when these ssNA regions are diminished.

It is widely appreciated that double stranded DNA (dsDNA) is subjected to strong and dynamic mechanical forces in cells. Under increasing tension B-DNA, the most stable double-stranded (ds) form of DNA, undergoes cooperative elongation into a mixture of S-DNA and single stranded DNA (ssDNA). Despite significant effort, the structure, energetics, kinetics and the biological role of S-DNA remains obscure. We here stretch 60 base pair (bp) dsDNA oligonucleotides with a variable number of tricyclic cytosine, tC, modifications using optical tweezers. We observe multiple fast cooperative and reversible two-state transitions between B-DNA and S-DNA. Notably, tC modifications increase the transition force, while reducing the transition extension and free energy due to progressively increasing fraying of the dsDNA ends. We quantify the average number of bps undergoing the B-to-S transition, as well as the free energies and rates. This allows us to reconstruct the B-to-S free energy profiles in absence of force. We conclude that S-DNA is an entirely force-induced state, and that the B-to-S transition is much faster than internal dsDNA melting. We hypothesize that S-DNA may have a role as a transient intermediate in, for example, molecular motor-induced local dsDNA strand separation.

Binuclear ruthenium complexes have been investigated for potential DNA-targeted therapeutic and diagnostic applications. Studies of DNA threading intercalation, in which DNA basepairs must be broken for intercalation, have revealed means of optimizing a model binuclear ruthenium complex to obtain reversible DNA-ligand assemblies with the desired properties of high affinity and slow kinetics. Here, we used single-molecule force spectroscopy to study a binuclear ruthenium complex with a longer semirigid linker relative to the model complex. Equilibrium results suggest a DNA affinity that is an order of magnitude higher than the parent binuclear ruthenium complex, likely due to a sterically relieved DNA threading intercalation mechanism. Notably, kinetics analysis shows that less DNA elongation is required for threading intercalation compared to the parent complex, and the association rate is two orders of magnitude faster. The ruthenium complex elongates the DNA duplex by ∼0.3 nm per bound ligand to reach the equilibrium intercalated state, with a significantly different energy landscape relative to the parent complex. Mechanical properties of the ligand-saturated DNA duplex show a higher persistence length, indicating that the longer semirigid linker provides enough molecular spacing to allow a single monomer to fully stack with basepairs, comparable to the monomeric parent ruthenium complex. The DNA basepairs in the equilibrium threading intercalated state are likely intact, and the ruthenium complex is shielded from the polar solution, providing measurable single-molecule confocal fluorescence signals. The obtained confocal fluorescence imaging of the bound dye confirms mostly uniform intercalation along the tethered DNA, consistent with other intercalators. The results of this study, along with previously examined ruthenium complex variants, illustrate tunable intercalation mechanisms guided by the rational design of therapeutic and diagnostic small molecules to target and modify the DNA duplex.

Noncanonical and damaged bases occur with significant frequency and recognizing them is an important step in many nuclear processes including DNA repair, homologous recombination, and transcription. Yet while both the pairing and stacking energies of the canonical bases are well known in theory and experiment, the energetics of noncanonical bases are not well understood. Incorporating noncanonical bases into DNA hairpins affords the opportunity to quantify the changes in stacking and pairing interactions during forced hairpin unfolding. Though the measurement accuracy of this technique is quite high, technical challenges are significant and the overall throughput is challenging. In this chapter, we present a method for measuring the pairing and stacking energies of such noncanonical base pairs by incorporating the hairpin defect in the lower stem and performing optical tweezers force-unfolding experiments. During unfolding, this stem frays apart before the upper half, enhancing detection and facilitating rapid and sensitive characterization of these defects.