Metal-dependent formate dehydrogenases reduce CO2 with high efficiency and selectivity, but are usually very oxygen sensitive. An exception is Desulfovibrio vulgaris W/Sec-FdhAB, which can be handled aerobically, but the basis for this oxygen tolerance was unknown. Here we show that FdhAB activity is controlled by a redox switch based on an allosteric disulfide bond. When this bond is closed, the enzyme is in an oxygen-tolerant resting state presenting almost no catalytic activity and very low formate affinity. Opening this bond triggers large conformational changes that propagate to the active site, resulting in high activity and high formate affinity, but also higher oxygen sensitivity. We present the structure of activated FdhAB and show that activity loss is associated with partial loss of the metal sulfido ligand. The redox switch mechanism is reversible in vivo and prevents enzyme reduction by physiological formate levels, conferring a fitness advantage during O2 exposure.

Arsenic contamination of groundwater is among one of the biggest health threats affecting millions of people in the world. There is an urgent need for efficient arsenic biosensors where the use of arsenic metabolizing enzymes can be explored. In this work, we have solved four crystal structures of arsenite oxidase (Aio) in complex with arsenic and antimony oxyanions and the structures determined correspond to intermediate states of the enzymatic mechanism. These structural data were complemented with density-functional theory calculations providing a unique view of the molybdenum active site at different time points that, together with mutagenesis data, enabled to clarify the enzymatic mechanism and the molecular determinants for the oxidation of As(III) to the less toxic As(V) species.Arsenic contamination of groundwater is among one of the biggest health threats affecting millions of people in the world. There is an urgent need for efficient arsenic biosensors where the use of arsenic metabolizing enzymes can be explored. In this work, we have solved four crystal structures of arsenite oxidase (Aio) in complex with arsenic and antimony oxyanions and the structures determined correspond to intermediate states of the enzymatic mechanism. These structural data were complemented with density-functional theory calculations providing a unique view of the molybdenum active site at different time points that, together with mutagenesis data, enabled to clarify the enzymatic mechanism and the molecular determinants for the oxidation of As(III) to the less toxic As(V) species.

A few ruthenium based metal carbonyl complexes, e.g. CORM-2 and CORM-3, have therapeutic activity attributed to their ability to deliver CO to biological targets. In this work, a series of related complexes with the formula [Ru(CO)(3)Cl2L] (L = DMSO (3), L-H3CSO(CH2)(2)CH(NH2)CO2H) (6a); D,L-H3CSO(CH2)(2)CH-(NH2)CO2H (6b); 3-NC5H4(CH2)(2)SO3.Na (7); 4-NC5H4(CH2)(2)SO3Na (8); PTA (9); DAPTA (10); H3CS-(CH2)(2)CH(OH) CO2H (11); CNCMe2CO2Me (12); CNCMeEtCO2Me (13); CN(c-C3H4)CO2Et) (14)) were designed, synthesized and studied. The effects of L on their stability, CO release profile, cytotoxicity and anti-inflammatory properties are described. The stability in aqueous solution depends on the nature of L as shown using HPLC and LC-MS studies. The isocyanide derivatives are the least stable complexes, and the S-bound methionine oxide derivative is the more stable one. The complexes do not release CO gas to the headspace, but release CO2 instead. X-ray diffraction of crystals of the model protein Hen Egg White Lysozyme soaked with 6b (4UWN) and 8 (4UWV) shows the addition of Ru-II(CO)(H2O)(4) at the His15 binding site. Soakings with 7 (4UWU) produced the metallacarboxylate [Ru(COOH)(CO)(H2O)(3)](+) bound to the His15 site. The aqueous chemistry of these complexes is governed by the water-gas shift reaction initiated with the nucleophilic attack of HO- on coordinated CO. DFT calculations show this addition to be essentially barrierless. The complexes have low cytotoxicity and low hemolytic indices. Following i.v. administration of CORM-3, the in vivo bio-distribution of CO differs from that obtained with CO inhalation or with heme oxygenase stimulation. A mechanism for CO transport and delivery from these complexes is proposed.

During the course of evolution, the cellulosome, one of Nature's most intricate multi-enzyme complexes, has been continuously fine-tuned to efficiently deconstruct recalcitrant carbohydrates. To facilitate the uptake of released sugars, anaerobic bacteria use highly ordered protein-protein interactions to recruit these nanomachines to the cell surface. Dockerin modules located within a non-catalytic macromolecular scaffold, whose primary role is to assemble cellulosomal enzymatic subunits, bind cohesin modules of cell envelope proteins, thereby anchoring the cellulosome onto the bacterial cell. Here we have elucidated the unique molecular mechanisms used by anaerobic bacteria for cellulosome cellular attachment. The structure and biochemical analysis of five cohesin-dockerin complexes revealed that cell surface dockerins contain two cohesin-binding interfaces, which can present different or identical specificities. In contrast to the current static model, we propose that dockerins utilize multivalent modes of cohesin recognition to recruit cellulosomes to the cell surface, a mechanism that maximises substrate access while facilitating complex assembly.

Two piano-stool iron(II) complexes bearing N-heterocyclic carbene ligands outfitted with acetamide- and amine-pendant arms [Cp*Fe(NHCR)(CO)I] {Cp* = η5-tetramethylcyclopentadienyl; R = CH2CONEt2 (3), (CH2)2NEt2 (4)}, have been prepared and fully characterized. Their catalytic activity in transfer hydrogenation (TH) of ketones using iPrOH as a hydrogen source and catalytic amounts of base (LiOtBu) has been explored, along with that of previously reported [CpFe(NHCR)(CO)I] {R = nBu (5), (CH2)2OH (6), Et (7), and (CH2)3OH (8)} complexes containing hydroxyl and nonfunctionalized alkyl arms. Complex 3 displayed the highest catalytic activity of the whole series 3–8, reaching a TOF50 value of 533 h–1. NMR monitoring of the stoichiometric reaction of 3 with LiOtBu, allowed the identification of a new species 3' containing a deprotonated amidate moiety, which has been fully characterized by 1H, 13C, and 15N NMR. Finally, a green protocol for the reduction of ketones through TH using glycerol as a hydrogen source, under microwave irradiation in the presence of catalytic amounts of 3 and base has been developed.

The Cellulosome is an intricate macromolecular protein complex that centralizes the cellulolytic efforts of many anaerobic microorganisms through the promotion of enzyme synergy and protein stability. The assembly of numerous carbohydrate processing enzymes into a macromolecular multiprotein structure results from the interaction of enzyme-borne dockerin modules with repeated cohesin modules present in noncatalytic scaffold proteins, termed scaffoldins. Cohesin–dockerin (Coh-Doc) modules are typically classified into different types, depending on structural conformation and cellulosome role. Thus, type I Coh-Doc complexes are usually responsible for enzyme integration into the cellulosome, while type II Coh-Doc complexes tether the cellulosome to the bacterial wall. In contrast to other known cellulosomes, cohesin types from Bacteroides cellulosolvens, a cellulosome-producing bacterium capable of utilizing cellulose and cellobiose as carbon sources, are reversed for all scaffoldins, i.e., the type II cohesins are located on the enzyme-integrating primary scaffoldin, whereas the type I cohesins are located on the anchoring scaffoldins. It has been previously shown that type I B. cellulosolvens interactions possess a dual-binding mode that adds flexibility to scaffoldin assembly. Herein, we report the structural mechanism of enzyme recruitment into B. cellulosolvens cellulosome and the identification of the molecular determinants of its type II cohesin–dockerin interactions. The results indicate that, unlike other type II complexes, these possess a dual-binding mode of interaction, akin to type I complexes. Therefore, the plasticity of dual-binding mode interactions seems to play a pivotal role in the assembly of B. cellulosolvens cellulosome, which is consistent with its unmatched complexity and size.

Arsenic is a widely distributed environmental toxin whose presence in drinking water poses a threat to >140 million people worldwide. The respiratory enzyme arsenite oxidase from various bacteria catalyses the oxidation of arsenite to arsenate and is being developed as a biosensor for arsenite. The arsenite oxidase from Rhizobium sp. str. NT-26 (a member of the Alphaproteobacteria) is a heterotetramer consisting of a large catalytic subunit (AioA), which contains a molybdenum centre and a 3Fe-4S cluster, and a small subunit (AioB) containing a Rieske 2Fe-2S cluster. Stopped-flow spectroscopy and isothermal titration calorimetry (ITC) have been used to better understand electron transfer through the redox-active centres of the enzyme, which is essential for biosensor development. Results show that oxidation of arsenite at the active site is extremely fast with a rate of >4000s−1 and reduction of the electron acceptor is rate-limiting. An AioB-F108A mutation results in increased activity with the artificial electron acceptor DCPIP and decreased activity with cytochrome c, which in the latter as demonstrated by ITC is not due to an effect on the protein-protein interaction but instead to an effect on electron transfer. These results provide further support that the AioB F108 is important in electron transfer between the Rieske subunit and cytochrome c and its absence in the arsenite oxidases from the Betaproteobacteria may explain the inability of these enzymes to use this electron acceptor.

Gram-positive bacteria homeostasis and antibiotic resistance mechanisms are dependent on the intricate architecture of the cell wall, where amidated peptidoglycan plays an important role. The amidation reaction is carried out by the bi-enzymatic complex MurT-GatD, for which biochemical and structural information is very scarce. In this work, we report the first crystal structure of the glutamine amidotransferase member of this complex, GatD from Staphylococcus aureus, at 1.85 Å resolution. A glutamine molecule is found close to the active site funnel, hydrogen-bonded to the conserved R128. In vitro functional studies using 1H-NMR spectroscopy showed that S. aureus MurT-GatD complex has glutaminase activity even in the absence of lipid II, the MurT substrate. In addition, we produced R128A, C94A and H189A mutants, which were totally inactive for glutamine deamidation, revealing their essential role in substrate sequestration and catalytic reaction. GatD from S. aureus and other pathogenic bacteria share high identity to enzymes involved in cobalamin biosynthesis, which can be grouped in a new sub-family of glutamine amidotransferases. Given the ubiquitous presence of GatD, these results provide significant insights into the molecular basis of the so far undisclosed amidation mechanism, contributing to the development of alternative therapeutics to fight infections.

Nucleotide-processing enzymes are key players in biological processes. They often operate through high substrate specificity for catalysis. How such specificity is achieved is unclear. Here, we dealt with this question by investigating all-α dimeric deoxyuridine triphosphate nucleotidohydrolases (dUTPases). Typically, these dUTPases hydrolyze either dUTP or deoxyuridine diphosphate (dUDP) substrates. However, the dUTPase enzyme DR2231 from Deinococcus radiodurans selectively hydrolyzes dUTP only, and not dUDP. By means of extended classical molecular dynamics simulations and quantum chemical calculations, we show that DR2231 achieves this specificity for dUTP via second-shell basic residues that, together with the two catalytic magnesium ions, contribute to properly orienting the γ-phosphate of dUTP in a prereactive state. This allows a nucleophilic water to be correctly placed and activated in order to perform substrate hydrolysis. We show that this enzymatic mechanism is not viable when dUDP is bound to DR2231. Importantly, in several other dUTPases capable of hydrolyzing either dUTP or dUDP, we detected that active site second-shell basic residues are more in number, anchoring the β-phosphate of the nucleotide substrate too, in contrast to what is observed in DR2231. Thus, strategically located basic second-shell residues mediate precise reactant positioning at the catalytic site, determining substrate specificity in dUTPases and possibly in other structurally similar nucleotide-processing metalloenzymes.Nucleotide-processing enzymes are key players in biological processes. They often operate through high substrate specificity for catalysis. How such specificity is achieved is unclear. Here, we dealt with this question by investigating all-α dimeric deoxyuridine triphosphate nucleotidohydrolases (dUTPases). Typically, these dUTPases hydrolyze either dUTP or deoxyuridine diphosphate (dUDP) substrates. However, the dUTPase enzyme DR2231 from Deinococcus radiodurans selectively hydrolyzes dUTP only, and not dUDP. By means of extended classical molecular dynamics simulations and quantum chemical calculations, we show that DR2231 achieves this specificity for dUTP via second-shell basic residues that, together with the two catalytic magnesium ions, contribute to properly orienting the γ-phosphate of dUTP in a prereactive state. This allows a nucleophilic water to be correctly placed and activated in order to perform substrate hydrolysis. We show that this enzymatic mechanism is not viable when dUDP is bound to DR2231. Importantly, in several other dUTPases capable of hydrolyzing either dUTP or dUDP, we detected that active site second-shell basic residues are more in number, anchoring the β-phosphate of the nucleotide substrate too, in contrast to what is observed in DR2231. Thus, strategically located basic second-shell residues mediate precise reactant positioning at the catalytic site, determining substrate specificity in dUTPases and possibly in other structurally similar nucleotide-processing metalloenzymes.

Understanding the specific molecular interactions between proteins and β1,3-1,4-mixed-linked d-glucans is fundamental to harvest the full biological and biotechnological potential of these carbohydrates and of proteins that specifically recognize them. The family 11 carbohydrate-binding module from Clostridium thermocellum (CtCBM11) is known for its binding preference for β1,3-1,4-mixed-linked over β1,4-linked glucans. Despite the growing industrial interest of this protein for the biotransformation of lignocellulosic biomass, the molecular determinants of its ligand specificity are not well defined. In this report, a combined approach of methodologies was used to unravel, at a molecular level, the ligand recognition of CtCBM11. The analysis of the interaction by carbohydrate microarrays and NMR and the crystal structures of CtCBM11 bound to β1,3-1,4-linked glucose oligosaccharides showed that both the chain length and the position of the β1,3-linkage are important for recognition, and identified the tetrasaccharide Glcβ1,4Glcβ1,4Glcβ1,3Glc sequence as a minimum epitope required for binding. The structural data, along with site-directed mutagenesis and ITC studies, demonstrated the specificity of CtCBM11 for the twisted conformation of β1,3-1,4-mixed-linked glucans. This is mediated by a conformation–selection mechanism of the ligand in the binding cleft through CH-π stacking and a hydrogen bonding network, which is dependent not only on ligand chain length, but also on the presence of a β1,3-linkage at the reducing end and at specific positions along the β1,4-linked glucan chain. The understanding of the detailed mechanism by which CtCBM11 can distinguish between linear and mixed-linked β-glucans strengthens its exploitation for the design of new biomolecules with improved capabilities and applications in health and agriculture. Database Structural data are available in the Protein Data Bank under the accession codes 6R3M and 6R31.