A single 16-base oligodeoxyribonucleotide was labeled at the 3′-end with fluorescein and at the 5′-end with x-rhodamine (R*oligo*F); the chromophores served as a donor/acceptor pair, respectively, for Förster resonance energy transfer. We exploited the striking differences in the steady-state emission spectra of the R*oligo*F as a single strand and in a duplex structure to signal hybridization in solution and to determine the kinetics of duplex formation as the probe bound to its oligomer complement and to its target sequence in M13mpl8(+) phage DNA. The binding followed second-order kinetics; in 0.18 M NaCl (pH 8) with 25% formamide, the rate constant for binding to the oligomer complement was 5.7 × 105 M-1 s-1, and that to M13mpl8(+) was 5.7 × 104 M-1 s-1. The source of the 10-fold decrease in the rate of binding to M13mpl8(+) was examined to differentiate between multiple nonproductive nucleation and rapid fluctuations in the structure around the target site. From simulations based on each model combined with associated experimental results, we concluded that the slower binding was due to rapid structural fluctuations around the target site, with an effective target concentration 0.1 of that of the total. Comparisons of total fluorescein emission derived from both steady-state and lifetime measurements suggest that the 5′-x-rhodamine induces a conformational change that affects the interaction at the 3′-end between the fluorescein and the polymer. The effects of salt on the fluorescence were complex. The static quenching of fluorescein in the single-labeled, single-stranded oligonucleotide did not change with NaCl (0–0.18 M), whereas there were marked changes in the double-labeled probe, showing that the conformational effects mediated by the 5′-x-rhodamine were salt dependent.
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