Enzyme Structure and Dynamics


Substrate selectivity and chemical specificity are the hallmarks of enzyme activity.   However, we still know relatively little about how the structure of an enzyme controls which substrate is bound, how substrate orientation is enforced in the active site, and what factors are responsible for chemical specificity in the reactions that are catalyzed by a given enzyme.  While crystallography can provide a three-dimensional structure that is invaluable in finding active sites and identifying catalytically important residues, these structures are primarily static in nature.  A photograph of a car engine, no matter how detailed, cannot provide much insight into what moves when the engine is running.   We need to be able to watch enzymes at work in four dimensions in order to really understand them, and time is the fourth dimension.   Nuclear magnetic resonance (NMR) can provide that fourth dimension, allowing us to watch changes in enzyme structure and dynamics in response to changes in substrate, cofactor and mutation.

Cytochrome P450- Nature’s blow-torch

Imagine you are a plumber fixing pipes in an old wooden house.  When it comes time to solder the pipes, you need to be very careful not to burn down the house with your torch.  Nature’s problem is similar.  We are surrounded by oxygen, a very reactive and corrosive substance.  The only thin
g standing between us a and spontaneous combustion is the fact that O2 exists normally in a triplet state (that is, with unpaired electrons) while most of the chemical bonds making up our bodies are singlet (all electrons paired), and don’t react easily with triplet species.  

When metabolic processes require a reaction with O2, it must be activated in order to accomplish the reaction.  Cytochromes P450 do this job efficiently, and catalyze many important reactions in our bodies, include steroid hormone and prostaglandin biosyntheses, as well as processing and metabolizing drugs.  For that matter, most antibiotics are biosynthesized at least in part by P450 enzymes, and many of the  microorganisms that we use to clean up toxic wastes and oil spills rely on P450 enzymes to do their work.  There are over 20,000 sequenced P450 genes in GenBank, covering all kingdoms and phyla of life.  Yet these diverse sequences, with thousands of different substrate/product combinations, all fold to a common, highly conserved three-dimensional structure.   

We are interested in how the P450 structure and dynamics are modulated in order to achieve the remarkable substrate selectivity and reaction specificity combined with high catalytic efficiency that many of these enzymes exhibit.  Our laboratory uses structural methods, especially NMR, combined with computational methods, bioinformatics, classical chemistry, molecular biology and enzymology to answer basic quest
ions about P450 function.   We recently discovered a hidden “spring” mechanism that adjusts the size of the active site in response to the size of bound substrate (Figure 1).  Many P450s require specific protein co-factors in order to turn over substrate. NMR methods have revealed that a series of complex changes take place in the P450 structure upon binding of the co-factor, and this has led us to propose a model for how these changes occur and why they are critical to enzymatic activity (Figures 2 and 3).   Currently, we are focusing on the specific residues that impart substrate specificity and product

Figure 1.  Left, hydrogen bonding patterns in the β3 sheet and K' helix, shown in relation to the heme and camphor in the CYP101A1 active site (from PDB entry 2CPP). Direction of movement of β3 sheet into active site upon conversion to of K' to 310 conformation is indicated by arrow. Right, details of the hydrogen bonding involving Gly 326, WAT 535, WAT 506 and Met 323.

selectivity on several P450 enzymes.

Figure 2.    Model for the complex between effector cofactor protein Pdx and P450 showing spatial distribution of Pdx-induced perturbations in the P450 molecule relative to the Pdx binding site.  Residues shown in blue are unperturbed, those in hot pink are perturbed by between 20 Hz and 160 Hz (i.e., they exhibit titratable or fast-exchange behavior) and those in green are perturbed by greater than 160 Hz (i.e., slow exchange behavior).



Figure 3.  Increased active site access from solvent in the trans conformer.

Left: The cis conformation showing Pro89 (in orange), Ile395 (light blue), Thr185 (dark blue) and bound camphor (magenta). Note the low solvent exposure of camphor. Right: The same view but in the trans conformation.  Camphor is more solvent exposed, indicating that the proposed substrate access channel is opened by the isomerization.

We are extending the methodology and expertise that we have developed to the important class of P450s that are involved in drug metabolism.  CYP3A4 is a human liver P450 that metabolizes over 70% of currently marketed pharmaceuticals, and defects in function of
CYP3A4 are important determinants of whether a person can tolerate a particular drug.  With the advent of personal genomes, understanding how particular mutations affect CYP3A4 function are becoming critical.  Because most mammalian P450s are membrane associated, they are difficult to work with using standard NMR methods.   We are experimenting with a novel membrane "sushi" called nanodiscs, developed by Steve Sligar at the University of Illinois, that solubilize membrane proteins in a native-like environment (Figure 4), so that we can apply high-resolution solution NMR methods to these important enzymes.

Figure 4.  CYP3A4 in nanodiscs.  For more info about the nanodisc technology see http://sligarlab.life.uiuc.edu/nanodisc.html


“Experimentally restrained molecular dynamics simulations for characterizing the open states of cytochrome P450cam." (E. K. Asciutto, M. Dang, S. S. Pochapsky, J. Madura, and T. C. Pochapsky) Biochemistry, 50, 1664–1671 (2011).

“Spring-loading the active site of cytochrome P450cam” (M. Dang, S. S. Pochapsky and T. C. Pochapsky) Metallomics 3, 339-343 (2011).

"Conformational plasticity and structure/function relationships in cytochromes P450" (T. C. Pochapsky, S. Kazanis and M. Dang), Antioxidants and Redox Signalling 13, 1273-1296 (2010).

"Redox-dependent dynamics in cytochrome P450cam" (S. S. Pochapsky, M. Dang, B. OuYang, A. K. Simorellis and T. C. Pochapsky), Biochemistry 48, 4254-4261 (2009).

"Structural and dynamic implications of an effector-induced backbone amide cis-trans isomerization in cytochrome P450cam" (E. K. Asciutto, J. D. Madura, S. S. Pochapsky, B. OuYang, T. C. Pochapsky) J. Mol. Biol. 388, 801-814 (2009).

"Solution NMR structure of putidaredoxin-cytochrome P450cam complex via a combined residual dipolar coupling-spin labeling approach suggests a role for Trp106 of putidaredoxin in complex formation" (W. Zhang, S. S. Pochapsky, T. C. Pochapsky and N. U. Jain)  J. Mol. Biol. 384, 349-363 (2008).

"A functional proline switch in cytochrome P450cam"   (B. OuYang, S. S. Pochapsky, M. Dang and T. C. Pochapsky) Cell Press Structure 16, 916-923 (2008).

"Hydrogen-deuterium exchange mass spectrometry for investigation of backbone dynamics of oxidized and reduced cytochrome P450cam" (Y. Hamuro, K. S. Molnar, S. J. Coales, B. OuYang, A. K. Simorellis and T. C. Pochapsky)  J. Inorg. Biochem. 102, 364-370 (2008).

“Specific effects of potassium ion binding on wild-type and L358P cytochrome P450cam

  1. B.OuYang, S. S. Pochapsky, G. M. Pagani, and T. C. Pochapsky) Biochemistry 45, 14379-14388 (2006).

“Comparison of the complexes formed by cytochrome P450cam with cytochrome b5 and putidaredoxin, two effectors of camphor hydroxylase activity” (L. Rui, S. S. Pochapsky & T. C. Pochapsky) Biochemistry 45 3887 – 3897 (2006).   

"Detection of a high-barrier conformational change in the active site of cytochrome P450cam upon binding of putidaredoxin" (J. Y. Wei, T. C. Pochapsky and S. S. Pochapsky), J. Am. Chem. Soc. 127, 6974-6976 (2005).  (Communication)

"A model for effector activity in a highly specific biological electron transfer complex: the cytochrome P450cam-putidaredoxin couple" (S. S. Pochapsky, T. C. Pochapsky and J. W. Wei), Biochemistry 42, 5649-5656 (2003).