The T7 RNA polymerase promoter has been proposed to contain two domains: the binding region upstream of position -5 is recognized through apparently traditional duplex contacts, while the catalytic domain downstream of position -5 is bound in a melted configuration. This model is tested by following polymerase binding to a series of synthetic oligonucleotides representing truncations of the consensus promoter sequence. The increase in the fluorescence anisotropy of a rhodamine dye linked to the upstream end of the promoter provides a very sensitive measure of enzyme binding in simple thermodynamic titrations, and allows the determination of both increases and decreases in the dissociation constant. The best fit value of K(d) = 4.0 nM for the native promoter is in good agreement with previous fluorescence and steady state measurements. Deletion of the downstream DNA up to position -1 or to position -5 leads to a fivefold increase in binding, while further sequential single-base deletions upstream result in 20 and 500-fold decreases in binding. These results indicate that the (duplex) region of the promoter upstream of and including position -5 is both necessary and sufficient for tight binding, and represents the core binding element of the promoter. We propose a model in which part of the upstream binding energy is used by T7 RNA polymerase to melt the downstream initiation region of the promoter. We also show that the presence of magnesium is necessary for optimal binding, but not for specific enzyme-promoter complex formation, and we propose that magnesium is not required for melting of the promoter.
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