Electrostatic-gated transport in chemically modified glass nanopore electrodes with orifice radii as small as 15 nm is reported. A single conical-shaped nanopore in glass, with a approximately 1 microm radius Pt disk located at the pore base, is prepared by etching the exposed surface of a glass-sealed Pt nanodisk. The electrochemical response of the nanopore electrode corresponds to diffusion of redox-active species through the nanopore orifice to the Pt microdisk. Silanization of the exterior glass surface with Cl(Me)(2)Si(CH(2))(3)CN and the interior pore surface with EtO(Me)(2)Si(CH(2))(3)NH(2) introduces pH-dependent ion selectivity at the pore orifice, a consequence of the electrostatic interactions between the redox ions and protonated surface amines. Nanopore electrodes with very small pore orifice radii (< approximately 50 nm) display anion permselectively at pH < 4, as demonstrated by electrochemical measurement of transport through the pore orifice. Ion selective transport vanishes at pH > 6 or when the pore radius is significantly larger than the Debye screening length, consistent with the observed ion selectivity resulting from electrostatic interactions. The ability to introduce different surface functionalities to the interior and exterior surfaces of glass nanopores is demonstrated using fluorescence microscopy to monitor the localized covalent attachment of 5- (and 6)-carboxytetramethylrhodamine succinimidyl ester to interior pore surfaces previously silanized with EtO(Me)(2)Si(CH(2))(3)NH(2).

The high affinity of avidin for biotin has made it useful for many bioanalytical applications involving the immobilization of proteins, vesicles, and other biomolecules to surfaces. To understand the formation and stability of the resulting biotin-avidin complex, it is useful to know the kinetics of the binding reaction, especially for situations where the complex is formed at a liquid-solid interface typically used in sensor or separation applications. In this work, a single-molecule fluorescence method is developed for measuring the kinetics and affinity constant for the binding of neutravidin, a deglycosylated variant of avidin, to surface-immobilized biotin. Biotin was immobilized using succinimidyl ester chemistry onto amine sites on glass surfaces. The surface density of biotin was controlled by the extreme dilution of 3-aminopropyltriethoxysilane into a monolayer of 2-cyanoethyltriethoxysilane. The resulting biotin binding sites are spaced apart by micrometer distances, and this avoids crowding effects and makes the resolution of single molecules possible. The binding and unbinding of individual tetramethylrhodamine-labeled neutravidin molecules is measured in situ by total-internal-reflection fluorescence (TIRF) microscopy imaging. Single-molecule detection and counting is readily achieved by this measurement, where quantitative control is established by determining the probabilities of false positive and negative events based on the intensity distributions of background and single-molecule spots and by comparing the bound molecule populations with the independently measured density of binding sites on the surface. The kinetics of binding and unbinding are evaluated by intermittent imaging and counting the number of bound neutravidin molecules versus time, following introduction of a neutravidin solution or its replacement by buffer over the low-density biotinylated surface. The neutravidin binding kinetics were found to be fast, essentially diffusion-controlled, while the stability of the complex and its dissociation rate appear to be influenced by the chemistry of biotin immobilization.

In recent years observations at the level of individual atoms and molecules became possible by micros-copy and spectroscopy. Imaging of single fluorescence molecules has been achieved but has so far been restricted to molecules in the immobile state. Here we provide methodology for visualization of the motion of individual fluorescent molecules. It is applied to imaging of the diffusional path of single molecules in a phospholipid membrane by using phospholip-ids carrying one rhodamine dye molecule. For this methodology , fluorescence microscopy was carried to a sensitivity so that single fluorescent molecules illuminated for only 5 ms were resolvable at a signal/noise ratio of 28. Repeated illuminations permitted direct observation of the diffusional motion of individual molecules with a positional accuracy of 30 nm. Such capability has fascinating potentials in bio-science-for example, to correlate biological functions of cell membranes with movements, spatial organization, and stoi-chiometries of individual components. The ultimate goal of high-sensitivity detection schemes is observation on the single molecule level. This came into reach by the invention of scanning probe microscopy (1, 2), which has since brought a wealth of new insights (3). Optical methods allowed for detection of single atoms (4). The effective light conversion in fluorescent molecules made it possible to detect single fluorophores in liquids by confocal fluorescence mi-croscopy (5-8) and to perform high-resolution spectroscopy of single dye molecules at low temperature (9-12). The first true imaging of single dye molecules by optical means was achieved by scanning near-field optical microscopy (13). This method is unique in reaching a spatial resolution of 14 nm, much below the optical diffraction limit but restricted in its application to immobile objects. Very recently, single fluorescence labeled myosin molecules on immobilized actin filaments were imaged by conventional microscopy and illumination times of seconds (14). It would be of interest for many applications, especially in bioscience, to extend microscopy to visualization of single fluorophores in motion. To our knowledge, such imaging has not been reported to date. Here we show that the motion of single dye molecules can be visualized by conventional fluo-rescence microscopy by extending the time resolution into the millisecond range. For this, we used epifluorescence micros-copy with argon-ion laser excitation and imaging onto a highly-sensitive liquid-nitrogen-cooled CCD-camera. Optical parts were carefully selected to achieve an efficiency for the detection of emitted fluorescence as high as 3%, while scattered light was blocked effectively. For demonstration of the potentials of observing individual mobile molecules we have chosen a fluorescence-labelled lipid in a fluid lipid membrane as a most appropriate system. It uniquely permitted to use results obtained at high surface densities of labelled lipid for The publication costs of this article were defrayed in part by p...

The density of surface-immobilized ligands or binding sites is an important issue for the development of sensors, array- or chip-based assays, and single-molecule detection methods. The goal of this research is to control the binding site density of reactive ligands on surfaces by diluting surface amine groups in self-assembled and cross-linked monolayers on glass prepared from solutions containing very low concentrations of (3-aminopropyl)triethoxysilane (APTES) and much higher concentrations of (2-cyanoethyl)triethoxysilane. The surface amine sites are suitable for attaching labels and ligands by reaction with succinimidyl ester reagents. Labeling the amine sites with fluorescent molecules and imaging the single molecules with fluorescence microscopy provides a means of determining the density of amine sites on the surface, which were incorporated into the self-assembled monolayer with micrometer spacings in proportion to the concentration of APTES in the synthesis. Biotin ligands were also bound to these surface amine sites using a succinimidyl ester linker, and the immobilized biotin was then reacted with either streptavidin-conjugated gold colloid particles or fluorescently labeled neutravidin. Imaging of these samples yields consistent amine and biotin site coverages, indicating that quantitative control and chemical conversion of binding sites can be achieved at very low (<10(-7)) fractions of a monolayer.

We quantify the adsorption and desorption of a monoclonal immunoglobulin-G antibody, rituxamab (RmAb), on silica capillary surfaces using electrospray-differential mobility analysis (ES-DMA). We first develop a theory to calculate coverages and desorption rate constants from the ES-DMA data for proteins adsorbing on glass capillaries used to electrospray protein solutions. This model is then used to study the adsorption of RmAb on a bare silica capillary surface. A concentration-independent coverage of ≈4.0 mg/m(2) is found for RmAb concentrations ranging from 0.01 to 0.1 mg/mL. A study of RmAb adsorption to bare silica as a function of pH shows maximum adsorption at its isoelectric point (pI of pH 8.5) consistent with literature. The desorption rate constants are determined to be ≈10(-5) s(-1), consistent with previously reported values, thus suggesting that shear forces in the capillary may not have a considerable effect on desorption. We anticipate that this study will allow ES-DMA to be used as a "label-free" tool to study adsorption of oligomeric and multicomponent protein systems onto fused silica as well as other surface modifications.