Specific placement of tryptophan in the catalytic sites of Escherichia coli F1-ATPase provides a direct probe of nucleotide binding: maximal ATP hydrolysis occurs with three sites occupied.

Residue beta Y331 of Escherichia coli F1-ATPase is known from previous affinity labeling, mutagenesis, and lin-benzo-ADP binding experiments to interact directly with the adenine moiety of substrates bound in catalytic sites. Here we mutagenized beta Y331 to tryptophan. Mutant cells grew well on succinate or limiting glucose; purified mutant F1 had kappa cat/Km and lin-benzo-ADP binding characteristics similar to wild type. Fluorescence from beta W331 residues exhibited a maximum at 349 nm, indicating a polar environment in unoccupied sites. ATP, ADP, or AMPPNP caused virtually complete quenching of beta W331 fluorescence, so that the fluorescence of mutant F1 with occupied catalytic sites resembled that of wild-type enzyme. Therefore the beta W331 fluorescence provided a direct probe of nucleotide binding to catalytic sites under true equilibrium conditions. We measured ATP binding and hydrolysis in parallel experiments and found that occupancy of one or two catalytic sites per F1 molecule did not yield significant rates of hydrolysis while occupancy of all three sites yielded Vmax rates. Km(ATP) was similar to Kd3, the Kd for ATP binding to the third catalytic site. We also measured AMPPNP and ADP binding parameters. For ADP, the "on" rate at the first catalytic site was much faster (> or = 5 x 10(5) M-1 s-1) than seen previously by centrifuge column procedures, although the Kd was not much changed. For AMPPNP, the "on" rate at the first site was 2 orders of magnitude less than for ADP or ATP, and the Kd was similar to that for ADP.