Biochemistry

Enzymes under the nanoscope

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Small-scale interactions of substrates with an enzyme's active site — over distances smaller than the length of a chemical bond — can make big differences to the enzyme's catalytic efficiency.

When Richard Feynman died in 1988, he left behind the following words on his blackboard: “What I cannot create, I do not understand.” His message certainly resonates with protein engineers. When it comes to making enzymes, we are clearly missing something, because artificial enzymes cannot yet be designed that match natural catalysts in efficiency. Reporting in the Journal of the American Chemical Society, Sigala et al.1 explain at least part of the reason why this is so. Tiny variations (on the scale of 10 picometres, where 1 picometre is 10−12 metres) in the binding interactions and molecular packing in an enzyme's active site can make a remarkable difference to the efficiency of enzymatic catalysis.

Any biochemistry textbook will tell you that enzymes catalyse reactions by binding their substrates' transition states — high-energy arrangements of atoms that form during reactions — more tightly than the ground states. This differential recognition lowers the energy barrier for reaction, and usually occurs because the transition state fits better into the active site than does the ground state, and/or because the active site stabilizes any charges in the transition state more than those in the ground state2.

Strong support for this idea comes from transition-state analogues (TSAs) — stable molecules designed to mimic the shapes and charges of transition states. TSAs are highly efficient inhibitors of enzyme catalysis because their tight binding to, and slow release from, enzymes' active sites blocks the turnover of native reactions3. Such molecules can even be used as templates to generate antibodies. Because these antibodies bind to TSAs, they should also bind to transition states for reactions modelled by the TSAs, thus catalysing those reactions. Such 'catalytic antibodies'4 are arguably the best models of enzymes that we have, but the reaction-rate accelerations of these proteins are still tens of billions of times smaller than those of many enzymes5,6.

Available tools for protein engineering clearly lack the subtle touch that is required to prepare effective designer enzymes. For example, site-directed mutagenesis (a method in which specific amino acids in proteins are replaced with others) is commonly used to investigate the roles of individual amino acids in catalysis. But the sizes of naturally occurring amino acids vary by discrete increments of at least one chemical-bond length (roughly 140 picometres), whereas breaking bonds in a transition state extend by only about 20 picometres, compared with the same bonds in the ground state. The modifications that we can make to active sites are therefore larger in scale than those that enzymes have evolved to detect. Further complications arise because, in a highly interconnected protein structure, a single amino-acid change introduced by site-directed mutagenesis can create all sorts of structural changes elsewhere in the enzyme.

Sigala et al.1 now report an approach for identifying the distance scale at which enzymes recognize the structural reorganization of substrates during reactions. Because transition states are, by definition, short-lived high-energy species that are not amenable to direct analysis, Sigala et al.1 had to investigate the effects on enzyme binding of structural variations in a TSA. They did this using a battery of modern analytical techniques — including high-resolution X-ray crystallography, nuclear magnetic resonance spectroscopy, quantum-mechanical calculations and TSA-binding measurements — that allowed them to resolve a difficult problem with unprecedented precision.

The enzyme chosen for study was ketosteroid isomerase (KSI), which catalyses the migration of a carbon–carbon double bond in a wide variety of ketosteroid substrates, by way of a negatively charged 'dienolate' intermediate (Fig. 1a, overleaf). The intermediate, and thus the transition state that leads to it, is stabilized by hydrogen bonding to hydroxyl (OH) groups in the side chains of two amino acids in the active site. These groups constitute an 'oxyanion hole' — a region of hydrogen-bonding groups capable of accommodating and stabilizing the negative charge that develops in the dienolate. Such oxyanion holes are held firmly in position by tight packing of local hydrophobic residues.

Figure 1: Enzyme-catalysed isomerization.
figure1

The enzyme ketosteroid isomerase (KSI) catalyses a reaction in which a carbon–carbon double bond in the substrate moves to a new position in the molecule. a, The side chain of an amino acid (red) in the active site triggers the reaction, which proceeds through a dienolate intermediate. Other side chains (green) stabilize the dienolate and the transition state that leads to it by forming hydrogen bonds (dashed lines) to its fully or partly negatively charged oxygen. These hydrogen bonds also bind the substrate and the product, albeit more weakly. Curly arrows indicate electron movement during the reaction. b, Sigala et al.1 investigate the control of KSI over the position of transition states during reactions, using phenolate ions as mimics of dienolates. Both the length of the carbon–oxygen bond and the electron density on the oxygen depend on the substituent X and on the bulkiness of the substituents R (R can be either hydrogen or fluorine).

The hydrogen bonds that stabilize the transition states in KSI also bind substrates in their ground states, but are presumed to 'tighten up' as the reaction proceeds. Sigala et al. monitored this tightening process using negatively charged phenolate ions as probes (Fig. 1b). Phenolates have a similar geometry and charge distribution to that of the dienolate, and bind to the active site of KSI using the same hydrogen bonds7. In KSI substrates, a carbon–oxygen double bond lengthens as the transition state forms, and the negative charge on the oxygen increases. Similarly, the length of an analogous carbon–oxygen bond in phenolates can be varied by changing a substituent on the phenolate; the electron density on the oxygen changes at the same time.

According to the accepted mechanism for KSI-catalysed reactions, increasing the electron density on the oxygen of a phenolate should strengthen (and shorten) the hydrogen bonds that bind the molecule to the active site, and so reinforce binding to the enzyme. Sigala et al.1 observe that this is indeed the case, but find that the pattern is disrupted if the phenolates are made slightly bulkier. When the hydrogen atoms attached to the carbons on either side of the oxygen are replaced by fluorine atoms (which are marginally larger and have higher electron density than hydrogens), increasing the electron density on the oxygen makes binding of the phenolate to the active site weaker, even though the hydrogen bonds should have been strengthened. This could be because the fluorine atoms start to clash (either electrostatically or physically) with the groups of the oxyanion hole as the hydrogen bonds try to become tighter, and thus shorter.

The crucial finding is that shortening of the hydrogen bonds by as little as 10 picometres is prevented by forces in and around the oxyanion hole, suggesting that the level of control exerted by the active site on the positions of its substrates operates on this stringently small scale. This result has wide-reaching implications: it defines experimentally the distance scale on which enzymes can distinguish geometric rearrangements of atoms, and determines the energetic consequences of this constraint. The picometre-precision of KSI also explains why protein engineering to produce enzymes that have new or altered functions has proved so difficult.

Sigala and colleagues' work brings one particular set of experimental tools to bear on a complex problem of fundamental importance, and will certainly concentrate the minds of those in the field. The same issue can and will be approached in other ways, and might well provide a range of answers that are specific to the system under investigation. We look forward to the development of a consensus. It will be interesting to see how other molecular probes can be used to map out the furnishings of active sites and to define and compare the distance scales for catalysis. Meanwhile, the belief that electrostatic and geometric complementarity of active sites and transition states is central to enzyme catalysis has become better defined. And, to accept Feynman's implicit challenge, what we understand, we might one day be able to create.

References

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Kirby, A., Hollfelder, F. Enzymes under the nanoscope. Nature 456, 45–47 (2008) doi:10.1038/456045a

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