Large-Scale Domain Conformational Change Is Coupled to the Activation of the Co–C Bond in the B₁₂-Dependent Enzyme Ornithine 4,5-Aminomutase: A Computational Study

We present here an energetic and atomistic description of how d-ornithine 4,5-aminomutase (OAM), an adenosylcobalamin (AdoCbl; coenzyme B₁₂)-dependent isomerase, employs a large-scale protein domain conformational change to orchestrate the homolytic rupture of the Co–C bond. Our results suggest that in going from the open form (catalytically inactive) to the closed form (catalytically active), the Rossmann domain of OAM effectively approaches the active site as a rigid body. It undergoes a combination of a ∼52° rotation and a ∼14 Å translation to bring AdoCblinitially positioned ∼25 Å awayinto the active-site cavity. This process is coupled to repositioning of the Ado moiety of AdoCbl from the eastern conformation to the northern conformation. Combined quantum mechanics and molecular mechanics calculations further indicate that in the open form, the protein environment does not impact significantly on the Co–C bond homolytic rupture, rendering it unusually stable, and thus catalytically inactive. Upon formation of the closed form, the Co–C bond is activated through the synergy of steric and electrostatic effects arising from tighter interactions with the surrounding enzyme. The more pronounced effect of the protein in the closed form gives rise to an elongated Co–C bond (by 0.03 Å), puckering of the ribose and increased “strain” energy on the Ado group and to a lesser extent the corrin ring. Our computational studies reveal novel strategies employed by AdoCbl-dependent enzymes in the control of radical catalysis

Proton-Coupled Electron Transfer and Adduct Configuration Are Important for C4a-Hydroperoxyflavin Formation and Stabilization in a Flavoenzyme

Determination of the mechanism of dioxygen activation by flavoenzymes remains one of the most challenging problems in flavoenzymology for which the underlying theoretical basis is not well understood. Here, the reaction of reduced flavin and dioxygen catalyzed by pyranose 2-oxidase (P2O), a flavoenzyme oxidase that is unique in its formation of C4a-hydroperoxyflavin, was investigated by density functional calculations, transient kinetics, and site-directed mutagenesis. Based on work from the 1970s–1980s, the current understanding of the dioxygen activation process in flavoenzymes is believed to involve electron transfer from flavin to dioxygen and subsequent proton transfer to form C4a-hydroperoxyflavin. Our findings suggest that the first step of the P2O reaction is a single electron transfer coupled with a proton transfer from the conserved residue, His548. In fact, proton transfer enhances the electron acceptor ability of dioxygen. The resulting ·OOH of the open-shell diradical pair is placed in an optimal position for the formation of C4a-hydroperoxyflavin. Furthermore, the C4a-hydroperoxyflavin is stabilized by the side chains of Thr169, His548, and Asn593 in a “face-on” configuration where it can undergo a unimolecular reaction to generate H₂O₂ and oxidized flavin. The computational results are consistent with kinetic studies of variant forms of P2O altered at residues Thr169, His548, and Asn593, and kinetic isotope effects and pH-dependence studies of the wild-type enzyme. In addition, the calculated energy barrier is in agreement with the experimental enthalpy barrier obtained from Eyring plots. This work revealed new insights into the reaction of reduced flavin with dioxygen, demonstrating that the positively charged residue (His548) plays a significant role in catalysis by providing a proton for a proton-coupled electron transfer in dioxygen activation. The interaction around the N5-position of the C4a-hydroperoxyflavin is important for dictating the stability of the intermediate

Mechanism of Action of Flavin-Dependent Halogenases

To rationally engineer the substrate scope and selectivity of flavin-dependent halogenases (FDHs), it is essential to first understand the reaction mechanism and substrate interactions in the active site. FDHs have long been known to achieve regioselectivity through an electrophilic aromatic substitution at C7 of the natural substrate Trp, but the precise role of a key active-site Lys residue remains ambiguous. Formation of hypochlorous acid (HOCl) at the cofactor-binding site is achieved by the direct reaction of molecular oxygen and a single chloride ion with reduced FAD and flavin hydroxide, respectively. HOCl is then guided 10 Å into the halogenation active site. Lys79, located in this site, has been proposed to direct HOCl toward Trp C7 through hydrogen bonding or a direct reaction with HOCl to form an −NH2Cl+ intermediate. Here, we present the most likely mechanism for halogenation based on molecular dynamics (MD) simulations and active-site density functional theory “cluster” models of FDH PrnA in complex with its native substrate l-tryptophan, hypochlorous acid, and the FAD cofactor. MD simulations with different protonation states for key active-site residues suggest that Lys79 directs HOCl through hydrogen bonding, which is confirmed by calculations of the reaction profiles for both proposed mechanisms

Scheme showing proposed step-wise conversion of Pchlide dimers to Chlide dimers. The I675* species only forms upon excitation of Pchlide-Chlide dimers.

<p>Scheme showing proposed step-wise conversion of Pchlide dimers to Chlide dimers. The I675* species only forms upon excitation of Pchlide-Chlide dimers.</p

Decay associated difference spectra (DADS) resulting from a global analysis of the pump-probe absorption spectroscopy data.

<p>Shown are DADS for Pchlide only (A), Chlide only (B), a sum of the ‘Pchlide only’ and ‘Chlide only’ data (C) and a mixture of 50% Pchlide and 50% Chlide, after photoexcitation at 460 nm. The data were fitted to a parallel model of independently decaying components as described in the <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0045642#s2" target="_blank">Materials and Methods</a> Section.</p

Residuals calculated after subtraction of the sum of the two component data from the time-resolved difference spectra for Pchlide and Chlide mixtures.

<p>The residuals calculated at 1 ns after subtraction of the sum of the ‘Pchlide only’ and ‘Chlide only’ data from the actual time-resolved difference spectra for all of the Pchlide and Chlide mixtures after photoexcitation at 450 nm (A), 460 nm (B) and 475 nm (C).</p

Pump-probe absorption spectroscopy of ‘Chlide only’ sample.

<p>Spectra are shown after photoexcitation with a laser pulse centred at ∼475 nm. The main panel shows transient absorption difference spectra at delay times of 2, 16, 516 and 2980 ps after excitation. The inset shows the respective kinetic transients at 675 nm (black circles) with a fit of the data to a single exponential (solid line).</p

Pump-probe absorption spectroscopy of samples containing a mixture of Pchlide and Chlide.

<p>Spectra are shown after photoexcitation with a laser pulse centred at ∼460 nm. The main panels show transient absorption difference spectra at delay times of 2, 16, 516 and 2980 ps after excitation for samples containing a mixture of 90% Pchlide and 10% Chlide (A), 50% Pchlide and 50% Chlide (B) and 10% Pchlide and 90% Chlide (C). The insets show the respective kinetic transients at 675 nm (black circles).</p

Pump-probe absorption spectroscopy of samples containing a mixture of 50% Pchlide and 50% Chlide in methanol after photoexcitation with a laser pulse centred at ∼460 nm.

<p>The main panel show transient absorption difference spectra at delay times of 2, 16, 516 and 2980 ps after excitation. The inset shows the residuals at 675 nm for a mixture of 50% Pchlide and 50% Chlide in methanol and H₂O after subtraction of the sum of the ‘Pchlide only’ and ‘Chlide only’ data from the actual time-resolved difference spectra.</p

Pump-probe absorption spectroscopy of POR-Pchlide-NADPH samples after photoexcitation with a laser pulse.

<p>The laser pulse was centred at ∼475 nm. Transient absorption difference spectra at delay times of 2, 17, 492 and 2992 ps after excitation are shown for the average of scans 1–3 (A), scans 8–10 (B) and scans 18–20 (C).</p