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.

Short double-stranded DNA is used in a variety of nanotechnological applications, and for many of them, it is important to know for which forces and which force loading rates the DNA duplex remains stable. In this work, we develop a theoretical model that describes the force-dependent dissociation rate for DNA duplexes tens of basepairs long under tension along their axes ("shear geometry"). Explicitly, we set up a three-state equilibrium model and apply the canonical transition state theory to calculate the kinetic rates for strand unpairing and the rupture-force distribution as a function of the separation velocity of the end-to-end distance. Theory is in excellent agreement with actual single-molecule force spectroscopy results and even allows for the prediction of the rupture-force distribution for a given DNA duplex sequence and separation velocity. We further show that for describing double-stranded DNA separation kinetics, our model is a significant refinement of the conventionally used Bell-Evans model.

Force-based ligand detection is a promising method to characterize molecular complexes label-free at physiological conditions. Because conventional implementations of this technique, e.g., based on atomic force microscopy or optical traps, are low-throughput and require extremely sensitive and sophisticated equipment, this approach has to date found only limited application. We present a low-cost, chip-based assay, which combines high-throughput force-based detection of dsDNA.ligand interactions with the ease of fluorescence detection. Within the comparative unbinding force assay, many duplicates of a target DNA duplex are probed against a defined reference DNA duplex each. The fractions of broken target and reference DNA duplexes are determined via fluorescence. With this assay, we investigated the DNA binding behavior of artificial pyrrole-imidazole polyamides. These small compounds can be programmed to target specific dsDNA sequences and distinguish between D- and L-DNA. We found that titration with polyamides specific for a binding motif, which is present in the target DNA duplex and not in the reference DNA duplex, reliably resulted in a shift toward larger fractions of broken reference bonds. From the concentration dependence nanomolar to picomolar dissociation constants of dsDNA.ligand complexes were determined, agreeing well with prior quantitative DNAase footprinting experiments. This finding corroborates that the forced unbinding of dsDNA in presence of a ligand is a nonequilibrium process that produces a snapshot of the equilibrium distribution between dsDNA and dsDNA.ligand complexes.

To locate its target site on DNA, a transcription factor (TF) must recognize its site amongst millions to billions of alternative sites on DNA. Studies suggested that TFs in order to facilitate their search process alternate between 3D diffusion in solution and 1D diffusion along DNA. The duration of such a search depends on the rate at which a TF slides along DNA and the frequency with which it alternates between 1D and 3D diffusion. We are interested in the 1D searching mechanism of p53, a transcription factor that functions as a tumor suppressor in human cells. We are using single-molecule techniques to observe diffusion of the fluorescently labeled p53 proteins along individual, stretched DNA molecules. In our previous studies, we determined the 1D diffusion coefficient of p53 protein. By measuring the 1D diffusion of the p53 protein as a function of ionic strength, we determined that the p53 protein maintains close contact with the DNA duplex and tracks the helical pitch. Current work involves the characterization of the role of the different protein domains in sliding. The C-terminus of p53 is suggested to be responsible for keeping the protein in contact with DNA by non-specifically interacting with the negatively charged backbone of DNA, while the core domain is suggested to be responsible for specifically binding the target site. We will present single-molecule data on the diffusional mobility along DNA of the C-terminal domain of p53, the p53 lacking its C-terminus, and the core domain of p53.

Der Molecular Force Assay ist eine kraftbasierte, hochgradig parallele Methode zur Charakterisierung biomolekularer Wechselwirkungen im Chipformat. Die Ergebnisse erlauben Rückschlüsse auf thermodynamische Größen und die Topologie der Bindungspotenziale.The Molecular Force Assay is a force-based, highly parallelized method in a chip format for the characterization of biomolecular interactions. Its results give information about thermodynamic constants, rates and the topology of the binding potential. Prinzip des Molecular Force Assayó Kraftspektroskopische Methoden wie das Rasterkraftmikroskop oder optische bzw. magnetische Fallen erlauben es, inter-und intramolekulare Wechselwirkungen, wie z. B. das Schmelzen von kurzer, doppelsträngiger DNA [1] oder die Bindungskraft zwischen Antikörper und Antigen [2], zu vermessen. Die zu untersuchende molekulare Bindung wird dazu mit einer bestimmten Kraft belastet und die Dehnung der Bindung bestimmt, sodass man eine Kraft-Abstands-Kurve erhält. Daraus lassen sich Informationen zur Abrisskraft, zur Entfaltung intramolekularer Proteindomänen oder zu den elastischen Eigenschaften der Biomoleküle bestimmen. Limitierend bei diesen Methoden ist die Kraftauflösung, da eine molekulare Größe mit einer mikrometerskaligen Blattfeder gemessen wird. Kleinere Federn werden weniger stark gedämpft, sodass das Rauschen vermindert wird und kleinere Kräfte aufgelöst werden können [3]. Allerdings können solche Federn aus technischen Gründen nicht beliebig klein hergestellt werden. Magnetische und optische Fallen ermöglichen die Detektion kleinerer Kräfte, können aber nur einen begrenzten Kraftbereich abdecken. Zusätzlich unterliegen die Wechselwirkungen zwischen Biomolekülen thermischen Fluktuationen, sodass nur über eine gute Statistik gesicherte Aussagen über Größen wie z. B. die Abrisskraft gemacht werden kön-nen.Um diese Einschränkungen zu umgehen, wurde der Molecular Force Assay (MFA) entwickelt. Hier wird die Feder durch eine zweite molekulare Bindung ersetzt, mit der die zu untersuchende Wechselwirkung verglichen wird, wodurch sehr kleine Unterschiede in der Stabilität der molekularen Bindungen detektiert werden können. Die zu untersuchende molekulare Bindung und der Referenzkomplex, eine kurze doppelsträngige DNA, werden in Serie auf einem Mikroskopierglas kovalent fixiert. Am oberen Ende des molekularen Komplexes ist ein Biotin angebunden, mit dem er an eine weiche Polydimethylsiloxan(PDMS)-Oberfläche, die mit Streptavidin funktionalisiert ist, binden kann (Abb. 1). Dieser PDMS-Stempel wird mit der Glasoberfläche vorsichtig in Kontakt gebracht, sodass die molekularen Bindungen über den Biotin-Streptavidin-Komplex zwischen den beiden Oberflächen eingespannt sind. Zieht man die weiche Oberfläche mithilfe eines Piezos zurück, wird eine Kraft in den molekularen Komplexen aufgebaut, die die Bindungen so lange belastet, bis die schwächere Wechselwirkung der beiden mit größerer Wahrscheinlichkeit reißt. Da an der Verbindungsstelle der beiden molekularen Komplexe ein Fluoreszenzfarbstoff ang...