Enzymes use substrate-binding energy both to promote ground-state association and to stabilize the reaction
transition state selectively. The
monomeric homing endonuclease I-AniI cleaves with high sequence specificity in the centre of a 20-base-pair (bp)
DNA target site, with the
amino (N)-terminal domain of the
enzyme making extensive binding interactions with the left (-) side of the target site and the similarly structured
carboxy (C)-terminal domain interacting with the right (+) side. Here we show that, despite the approximate twofold symmetry of the enzyme-DNA complex, there is almost complete segregation of interactions responsible for
substrate binding to the (-) side of the interface and interactions responsible for transition-state stabilization to the (+) side. Although single
base-pair substitutions throughout the entire
DNA target site reduce
catalytic efficiency,
mutations in the (-)
DNA half-site almost exclusively increase the
dissociation constant (K(D)) and the
Michaelis constant under single-turnover conditions (K(M)*), and those in the (+) half-site primarily decrease the
turnover number (k(cat)*). The reduction of activity produced by
mutations on the (-) side, but not
mutations on the (+) side, can be suppressed by
tethering the
substrate to the
endonuclease displayed on the surface of
yeast. This dramatic asymmetry in the use of enzyme-substrate binding energy for
catalysis has direct relevance to the redesign of
endonucleases to cleave genomic target sites for
gene therapy and other applications. Computationally redesigned
enzymes that achieve new specificities on the (-) side do so by modulating K(M)*, whereas redesigns with altered specificities on the (+) side modulate k(cat)*. Our results illustrate how classical
enzymology and modern
protein design can each
inform the other.
