DNA methylation plays an essential role in transcriptional control of organismal development in epigenetics, from turning off a specific gene to inactivation of entire chromosomes. While the biological function of DNA methylation is becoming increasingly clear, the mechanism of methylation-induced gene regulation is still poorly understood. Through single-molecule force experiments and simulation we investigated the effects of methylation on strand separation of DNA, a crucial step in gene expression. Molecular force assay and single-molecule force spectroscopy revealed a strong methylation dependence of strand separation. Methylation is observed to either inhibit or facilitate strand separation, depending on methylation level and sequence context. Molecular dynamics simulations provided a detailed view of methylation effects on strand separation, suggesting the underlying physical mechanism. According to our study, methylation in epigenetics may regulate gene expression not only through mechanisms already known but also through changing mechanical properties of DNA.

While nanophotonic devices are unfolding their potential for single-molecule fluorescence studies, metallic quenching and steric hindrance, occurring within these structures, raise the desire for site-specific immobilization of the molecule of interest. Here, we refine the single-molecule cut-and-paste technique by optical superresolution routines to immobilize single fluorescent molecules in the center of nanoapertures. By comparing their fluorescence lifetime and intensity to stochastically immobilized fluorophores, we characterize the electrodynamic environment in these nanoapertures and proof the nanometer precision of our loading method.

Without prior signal amplification, small molecules are difficult to detect by current label-free biochip approaches. In the present study, we developed a label-free capture biochip based on the comparative measurement of unbinding forces allowing for direct detection of small-molecule-aptamer interactions. The principle of this assay relies on increased unbinding forces of bipartite aptamers due to complex formation with their cognate ligands. The bipartite aptamers are immobilized on glass support via short DNA duplexes that serve as references to which unbinding forces can be compared. In a simple model system, adenosine is captured from solution by an adenosine-selective aptamer. Linking the molecular chains, each consisting of a short DNA reference duplex and a bipartite aptamer, between glass and a poly(dimethylsiloxane) (PDMS) surface and subsequently separating the surfaces compares the unbinding forces of the two bonds directly. Fluorescence readout allows for quantification of the fractions of broken aptamer and broken reference bonds. The presence of micromolar adenosine concentrations reliably resulted in a shift toward larger fractions of broken reference bonds. Because of the force-based design, the interactions between the bipartite aptamer and the target, rather than the presence of the target, are detected and no washing step disturbing the equilibrium state prior to probing and no reporter aptamer or antibody is required. The assay exhibits excellent selectivity against other nucleotides and detects adenosine in the presence of a complex molecular background. Multiplexing was demonstrated by performing whole titration experiments on a single chip revealing an effective half-maximal concentration of 124.8 microM agreeing well with literature values.

Cytosine hydroxymethylation is an epigenetic control factor in higher organisms. New discoveries of the biological roles of hydroxymethylation serve to raise questions about how this epigenetic modification exerts its functions and how organisms discriminate cytosine hydroxymethylation from methylation. Here, we report investigations that reveal an effect of cytosine hydroxymethylation on mechanical properties of DNA under load. The findings are based on molecular force assay measurements and steered molecular dynamics simulations. Molecular force assay experiments identified significant effects of hydroxymethylation on stretching-induced strand separation; the underlying physical mechanism has been revealed by steered molecular dynamics simulations. We find that hydroxymethylation can either upregulate or downregulate DNA's strand separation propensity, suggesting that hydroxymethylation can control gene expression by facilitating or obstructing the action of transcription machinery or the access to chromosomal DNA.