CO and CN? syntheses by [FeFe]-hydrogenase maturase HydG are catalytically differentiated events

The synthesis and assembly of the active site [FeFe] unit of [FeFe]-hydrogenases require at least three maturases. The radical S-adenosyl-l-methionine HydG, the best characterized of these proteins, is responsible for the synthesis of the hydrogenase CO and CN⁻ ligands from tyrosine-derived dehydroglycine (DHG). We speculated that CN⁻ and the CO precursor ⁻:CO₂H may be generated through an elimination reaction. We tested this hypothesis with both wild type and HydG variants defective in second iron-sulfur cluster coordination by measuring the in vitro production of CO, CN⁻, and ⁻:CO₂H-derived formate. We indeed observed formate production under these conditions. We conclude that HydG is a multifunctional enzyme that produces DHG, CN⁻, and CO at three well-differentiated catalytic sites. We also speculate that homocysteine, cysteine, or a related ligand could be involved in Fe(CO)_x(CN)_y transfer to the HydF carrier/scaffold.Many microorganisms can either oxidize molecular hydrogen or reduce protons according to the reaction H₂ = 2H⁺ + 2 e⁻. The enzymes that catalyze this reaction fall into two phylogenetically unrelated groups, the [NiFe]- and [FeFe]-hydrogenases (1, 2). Initial crystallographic studies of the [FeFe]-hydrogenases from Clostridium pasteuranium (3) and Desulfovibrio desulfuricans (4) showed that the active site is composed of a conventional [4Fe-4S] cubane connected by a cysteine thiolate to a binuclear FeFe unit, in which each iron ion is terminally coordinated by one CN⁻ ligand and one CO ligand and by a third CO molecule that bridges the two metals (5). Unexpectedly, we also found that a small molecule first postulated (6), and now indirectly confirmed (7), to be dithiomethylamine (DTMA) bridges the two Fe ions (Fig. 1).Open in a separate windowFig. 1.[FeFe]-hydrogenase H-cluster from D. desulfuricans (6). Only the cysteine residue bridging the [4Fe-4S] and [FeFe] subsites is depicted as Cys. The cysteine ligands of the [4Fe-4S] cluster are shown as straight lines.The [4Fe-4S] cubane bridged to the binuclear [FeFe] unit has been collectively called the H-cluster (1). Work from several laboratories has shown that the maturation of the [FeFe] center requires at least three protein maturases: HydF that has GTPase activity and appears to be both a [FeFe] center scaffold and carrier (8, 9), HydG that synthesizes CO and CN⁻ from tyrosine (10–13), and HydE that, by elimination, should be involved in the synthesis of the DTMA bridge (14, 15). Both HydE and HydG are members of the large radical S-adenosyl-l-methionine (SAM) protein family (16, 17). With the recent reports of HydG crystal structures from Carboxydothermus hydrogenoformans (Ch) by us (18) and from Thermoanaerobacter italicus (Ti) by Dinis et al. (19), X-ray models are now available for the three maturases (20, 21); however, unambiguous structure-function relationships have been proposed only in the case of HydG. Indeed, site-directed mutational studies have shown that CO and CN⁻ syntheses are affected by either the deletion of the maturase C-terminal region, where a second iron-sulfur cluster binds (22), or Cys-to-Ser mutations in its corresponding CxxCx₂₂C binding motif (10, 13). In addition, it has been shown that HydG synthesizes Fe(CO)_x(CN)_y precursors (x = 1 or 2; y = 1) of the [FeFe] catalytic unit (23). The two HydG crystal structures are very similar at the SAM and [4Fe-4S] cluster-containing (β/α)₈ TIM-like barrel, common to several radical SAM proteins (16) (Fig. 2). Conversely, there are significant differences in the composition of the extra C-terminal second (s) iron-sulfur cluster. In our crystals, ChHydG lacks this center (18), whereas TiHydG coordinates a [4Fe-4S]_s cluster in one of the two molecules of the asymmetric unit and a second center with a fifth iron in the other molecule (19). This fifth iron has been described as being bound by His265, a putative alanine molecule and a sulfide bridge to a [4Fe-4S] unit coordinated by the CxxCx₂₂C motif. Two water molecules complete the octahedral Fe coordination (19). Here the different second cluster structures are collectively called [FeS]_s.Open in a separate windowFig. 2.Structures of (A) Ch HydG depicting tunnel I, the SAM cofactor, and [4Fe-4S] cluster (Top Right), the tyrosine active site cavity (Top Center), tunnel II, and the Cl⁻ binding cavity (Bottom Center) (18) and (B) Ti HydG with its additional second iron-sulfur cluster (19).     

Role of protein dynamics in ion selectivity and allosteric coupling in the NaK channel

Flux-dependent inactivation that arises from functional coupling between the inner gate and the selectivity filter is widespread in ion channels. The structural basis of this coupling has only been well characterized in KcsA. Here we present NMR data demonstrating structural and dynamic coupling between the selectivity filter and intracellular constriction point in the bacterial nonselective cation channel, NaK. This transmembrane allosteric communication must be structurally different from KcsA because the NaK selectivity filter does not collapse under low-cation conditions. Comparison of NMR spectra of the nonselective NaK and potassium-selective NaK2K indicates that the number of ion binding sites in the selectivity filter shifts the equilibrium distribution of structural states throughout the channel. This finding was unexpected given the nearly identical crystal structure of NaK and NaK2K outside the immediate vicinity of the selectivity filter. Our results highlight the tight structural and dynamic coupling between the selectivity filter and the channel scaffold, which has significant implications for channel function. NaK offers a distinct model to study the physiologically essential connection between ion conduction and channel gating.Ion conduction through the pore domain of cation channels is regulated by two gates: an inner gate at the bundle crossing of the pore-lining transmembrane helices and an outer gate located at the selectivity filter (Fig. 1 B and C). These two gates are functionally coupled as demonstrated by C-type inactivation, in which channel opening triggers loss of conduction at the selectivity filter (1–4). A structural model for C-type inactivation has been developed for KcsA, with selectivity filter collapse occurring upon channel opening (4–10). In the reverse pathway, inactivation of the selectivity filter has been linked to changes at the inner gate (5–14). However, flux-dependent inactivation occurs in Na⁺ and Ca²⁺ channels as well and would likely require a structurally different mechanism to explain coupling between the selectivity filter and inner gate (7, 13–18).Open in a separate windowFig. 1.Crystal structures of the nonselective cation channel NaK and the potassium-selective NaK2K mutant show structural changes restricted to the area of the selectivity filter. Alignment of the WT NaK (gray; PDB 3E8H) and NaK2K (light blue; PDB 3OUF) selectivity filters shows a KcsA-like four-ion-binding-site selectivity filter is created by the NaK2K mutations (D66Y and N68D) (A), but no structural changes occur outside the vicinity of the selectivity filter (B). (C) Full-length NaK (green; PDB 2AHZ) represents a closed conformation. Alignment of this structure with NaK (gray) highlights the changes in the M2 hinge (arrow), hydrophobic cluster (residues F24, F28, and F94 shown as sticks), and constriction point (arrow; residue Q103 shown as sticks) upon channel opening. Two (A) or three monomers (B and C) from the tetramer are shown for clarity.This study provides experimental evidence of structural and dynamic coupling between the inner gate and selectivity filter in the NaK channel, a nonselective cation channel from Bacillus cereus (19). These results were entirely unexpected given the available high-resolution crystal structures (20, 21). The NaK channel has the same basic pore architecture as K⁺ channels (Fig. 1 B and C) and has become a second model system for investigating ion selectivity and gating due to its distinct selectivity filter sequence (₆₃TVGDGN₆₈) and structure (19–23). Most strikingly, there are only two ion binding sites in the selectivity filter of the nonselective NaK channel (Fig. 1A) (21, 24). However, mutation of two residues in the selectivity filter sequence converts the NaK selectivity filter to the canonical KcsA sequence (₆₃TVGYGD₆₈; Fig. 1 A and B), leading to K⁺ selectivity and a KcsA-like selectivity filter structure with four ion binding sites (21, 23). This K⁺-selective mutant of NaK is called NaK2K. Outside of the immediate vicinity of the two mutations in the selectivity filter, high-resolution crystal structures of NaK and NaK2K are essentially identical (Fig. 1B) with an all-atom rmsd of only 0.24 Å.NaK offers a distinct model to study the physiologically essential connection between ion conduction and channel gating because there is no evidence for any collapse or structural change in the selectivity filter. The NaK selectivity filter structure is identical in Na⁺ or K⁺ (22) and even in low-ion conditions (25), consistent with its nonselective behavior. Even the selective NaK2K filter appears structurally stable in all available crystal structures (25). Here we use NMR spectroscopy to study bicelle-solubilized NaK. Surprisingly, we find significant differences in the NMR spectra of NaK and NaK2K that extend throughout the protein and are not localized to the selectivity filter region. This, combined with NMR dynamics studies of NaK, suggests a dynamic pathway for transmembrane coupling between the inner gate and selectivity filter of NaK.     

Heterogeneity of functional groups in a metal–organic framework displays magic number ratios

Multiple organic functionalities can now be apportioned into nanoscale domains within a metal-coordinated framework, posing the following question: how do we control the resulting combination of “heterogeneity and order”? Here, we report the creation of a metal–organic framework, MOF-2000, whose two component types are incorporated in a 2:1 ratio, even when the ratio of component types in the starting solution is varied by an order of magnitude. Statistical mechanical modeling suggests that this robust 2:1 ratio has a nonequilibrium origin, resulting from kinetic trapping of component types during framework growth. Our simulations show how other “magic number” ratios of components can be obtained by modulating the topology of a framework and the noncovalent interactions between component types, a finding that may aid the rational design of functional multicomponent materials.The assembly of multiple types of component offers a potential route to the precise control of component heterogeneity within ordered 3D frameworks. Metal–organic frameworks (MOFs) (1–5) possessing well-defined connectivities (6, 7) and tunable pore sizes (8–10) can assemble from a variety of building blocks (11–17). Recently, a MOF harboring two components distributed in a heterogeneous fashion on an ordered framework was demonstrated (1, 2). There exists no framework, however, whose component heterogeneity remains controlled in the face of changes of environment. Here, we report the creation of a material with exactly this property. MOF-2000 is assembled from two types of organic struts, called L_r and L_b (Fig. 1A). These struts have identical rigid backbones but bear either a crown ether (L_r) or [2]catenane (L_b) side chain attached at their center. X-ray diffraction of MOF-2000 single crystals revealed that struts form a twofold interpenetrated cubic framework of pcu-c topology (Fig. 1A, Right, and SI Appendix, section S1.3; structure available in Dataset S1). As a result of optical investigations, we confirmed that the two struts are distributed in the crystalline framework in an isotropic manner (Fig. 1B), suggesting that the two components are not distributed in a simple periodic way throughout the framework (because such arrangements would give rise to optical anisotropy; SI Appendix, section S1.5).Open in a separate windowFig. 1.MOF-2000 is a periodic framework harboring controlled component heterogeneity. (A) Chemical structures of organic struts H₂L_r and H₂L_b⁴⁺ incorporated in MOF-2000 (Methods Summary), and the MOF-2000 crystal structure [key: carbon, black; oxygen, red; Zn₄O(CO₂)₆ polyhedra, blue; all hydrogen atoms have been omitted for clarity]. (B) Optical images of MOF-2000 single crystal: without polarizer (up) and in between crossed polarizers (down). The lack of birefringence indicates that MOF-2000 is optically isotropic. The edge of the crystal is about 0.1 mm. (C) As determined by ¹H-NMR and powder X-ray diffraction measurements (SI Appendix, sections S1.3 and S1.4), MOF-2000 consists of a twofold interpenetrated crystalline framework (of topology pcu-c) of 2:1 L_r:L_b ratio (estimated 5% error), for a wide range of solution ratios (shaded region). (D) MOF-2000 therefore combines the regularity of a covalent framework with controlled irregularity of its components.Even though the arrangement of the two components in MOF-2000 is indiscernible by X-ray crystallography, presumably as a result of the positional disorder of the two strut types within the framework, and the rotational (11, 18) and conformational (11) disorder of the side chains, the presence of the two organic struts can be clearly determined by ¹H-NMR (SI Appendix, section S1.4). Strikingly, MOF-2000 displays a 2:1 ratio of L_r and L_b struts, even when the ratios of components in the parent solution are varied by over an order of magnitude (Fig. 1C). This feature makes MOF-2000 unique among multicomponent extended frameworks, whose struts are usually incorporated in ratios determined largely by initial solution conditions (1, 19). Thus, MOF-2000 incorporates environmentally robust component heterogeneity within a regular covalent framework (Fig. 1D).The robust 2:1 composition does not have an obvious thermodynamic origin, in contrast to common organic cocrystals or ionic systems such as CsCl (20). The latter possess robust compositions as a consequence of charge complementarity. In MOF-2000, the L_b strut bears a charge of +4 (neutralized in the assembly by counterions), whereas the L_r strut is uncharged. Thus, charge complementarity alone does not suggest a 2:1 component ratio. Simple packing arguments suggest that it is reasonable that L_b is not the majority species within the framework (because it is bulky), but likewise do not offer a simple explanation for the 2:1 ratio.     

Tuning a riboswitch response through structural extension of a pseudoknot

Structural and dynamic features of RNA folding landscapes represent critical aspects of RNA function in the cell and are particularly central to riboswitch-mediated control of gene expression. Here, using single-molecule fluorescence energy transfer imaging, we explore the folding dynamics of the preQ₁ class II riboswitch, an upstream mRNA element that regulates downstream encoded modification enzymes of queuosine biosynthesis. For reasons that are not presently understood, the classical pseudoknot fold of this system harbors an extra stem–loop structure within its 3′-terminal region immediately upstream of the Shine–Dalgarno sequence that contributes to formation of the ligand-bound state. By imaging ligand-dependent preQ₁ riboswitch folding from multiple structural perspectives, we reveal that the extra stem–loop strongly influences pseudoknot dynamics in a manner that decreases its propensity to spontaneously fold and increases its responsiveness to ligand binding. We conclude that the extra stem–loop sensitizes this RNA to broaden the dynamic range of the ON/OFF regulatory switch.A variety of small metabolites have been found to regulate gene expression in bacteria, fungi, and plants via direct interactions with distinct mRNA folds (1–4). In this form of regulation, the target mRNA typically undergoes a structural change in response to metabolite binding (5–9). These mRNA elements have thus been termed “riboswitches” and generally include both a metabolite-sensitive aptamer subdomain and an expression platform. For riboswitches that regulate the process of translation, the expression platform minimally consists of a ribosomal recognition site [Shine–Dalgarno (SD)]. In the simplest form, the SD sequence overlaps with the metabolite-sensitive aptamer domain at its downstream end. Representative examples include the S-adenosylmethionine class II (SAM-II) (10) and the S-adenosylhomocysteine (SAH) riboswitches (11, 12), as well as prequeuosine class I (preQ₁-I) and II (preQ₁-II) riboswitches (13, 14). The secondary structures of these four short RNA families contain a pseudoknot fold that is central to their gene regulation capacity. Although the SAM-II and preQ₁-I riboswitches fold into classical pseudoknots (15, 16), the conformations of the SAH (17) and preQ₁-II counterparts are more complex and include a structural extension that contributes to the pseudoknot architecture (14). Importantly, the impact and evolutionary significance of these “extra” stem–loop elements on the function of the SAH and preQ₁-II riboswitches remain unclear.PreQ₁ riboswitches interact with the bacterial metabolite 7-aminomethyl-7-deazaguanine (preQ₁), a precursor molecule in the biosynthetic pathway of queuosine, a modified base encountered at the wobble position of some transfer RNAs (14). The general biological significance of studying the preQ₁-II system stems from the fact that this gene-regulatory element is found almost exclusively in the Streptococcaceae bacterial family. Moreover, the preQ₁ metabolite is not generated in humans and has to be acquired from the environment (14). Correspondingly, the preQ₁-II riboswitch represents a putative target for antibiotic intervention. Although preQ₁ class I (preQ₁-I) riboswitches have been extensively investigated (18–28), preQ₁ class II (preQ₁-II) riboswitches have been largely overlooked despite the fact that a different mode of ligand binding has been postulated (14).The consensus sequence and the secondary structure model for the preQ₁-II motif (COG4708 RNA) (Fig. 1A) comprise ∼80–100 nt (14). The minimal Streptococcus pneumoniae R6 aptamer domain sequence binds preQ₁ with submicromolar affinity and consists of an RNA segment forming two stem–loops, P2 and P4, and a pseudoknot P3 (Fig. 1B). In-line probing studies suggest that the putative SD box (AGGAGA; Fig. 1) is sequestered by pseudoknot formation, which results in translational-dependent gene regulation of the downstream gene (14).Open in a separate windowFig. 1.PreQ₁ class II riboswitch. (A) Chemical structure of 7-aminomethyl-7-deazaguanosine (preQ₁); consensus sequence and secondary structure model for the COG4708 RNA motif (adapted from reference 14). Nucleoside presence and identity as indicated. (B) S. pneumoniae R6 preQ₁-II RNA aptamer investigated in this study. (C) Schematics of an H-type pseudoknot with generally used nomenclature for comparison.Here, we investigated folding and ligand recognition of the S. pneumoniae R6 preQ₁-II riboswitch, using complementary chemical, biochemical, and biophysical methods including selective 2′-hydroxyl acylation analyzed by primer extension (SHAPE), mutational analysis experiments, 2-aminopurine fluorescence, and single-molecule fluorescence resonance energy transfer (smFRET) imaging. In so doing, we explored the structural and functional impact of the additional stem–loop element in the context of its otherwise “classical” H-type pseudoknot fold (29–32) (Fig. 1C). Our results reveal that the unique 3′-stem–loop element in the preQ₁-II riboswitch contributes to the process of SD sequestration, and thus the regulation of gene expression, by modulating both its intrinsic dynamics and its responsiveness to ligand binding.     

Detection of an intermediary,protonated water cluster in photosynthetic oxygen evolution

In photosynthesis, photosystem II evolves oxygen from water by the accumulation of photooxidizing equivalents at the oxygen-evolving complex (OEC). The OEC is a Mn₄CaO₅ cluster, and its sequentially oxidized states are termed the S_n states. The dark-stable state is S₁, and oxygen is released during the transition from S₃ to S₀. In this study, a laser flash induces the S₁ to S₂ transition, which corresponds to the oxidation of Mn(III) to Mn(IV). A broad infrared band, at 2,880 cm⁻¹, is produced during this transition. Experiments using ammonia and ²H₂O assign this band to a cationic cluster of internal water molecules, termed “W₅⁺.” Observation of the W₅⁺ band is dependent on the presence of calcium, and flash dependence is observed. These data provide evidence that manganese oxidation during the S₁ to S₂ transition results in a coupled proton transfer to a substrate-containing, internal water cluster in the OEC hydrogen-bonded network.Internal proton transfer reactions play important catalytic roles in many integral membrane proteins. In these enzymes, including bacteriorhodopsin, light-driven or redox-coupled proton-transfer reactions lead to the production of a transmembrane, electrochemical gradient. Amino acid side chains often participate in the acid/base chemistry that occurs in proton-transfer pathways. However, internal bound water clusters can also play essential roles as proton donors or acceptors (reviewed in ref. 1).In photosystem II (PSII), proton transfer contributes to the generation of a transmembrane potential, and chemical protons are released from the substrate, water, during the light-driven reactions that produce molecular oxygen (2). PSII is a complex membrane protein consisting of both integral, membrane-spanning subunits and extrinsic subunits (3). A monomeric unit of PSII consists of at least 20 distinct protein subunits, which are composed of 17 integral subunits and 3 extrinsic polypeptides (4, 5). The primary subunits that make up the reaction center and bind most of the redox-active cofactors are D1, D2, CP43, and CP47. The light-induced electron transfer pathway in the reaction center involves the dimeric chlorophyll (chl) donor, P₆₈₀, and accessory chl molecules. One light-induced charge separation oxidizes the primary donor, P₆₈₀, and reduces a bound plastoquinone acceptor, Q_A. P₆₈₀⁺ oxidizes a tyrosine residue, YZ, Y161 of the D1 polypeptide, which is a powerful oxidant. YZ• oxidizes the oxygen-evolving complex (OEC) on each photoinduced charge separation (reviewed in ref. 6).The OEC is a Mn₄CaO₅ cluster (Fig. 1 A, Inset) (5). Oxygen release from the OEC fluctuates with period four (7). The OEC cycles through five sequentially oxidized states, called the S_n states. A single flash given to a dark-adapted sample (S₁ state) generates the S₂ state (Fig. 1A), which corresponds to the oxidation of Mn(III) to Mn(IV) (8). Subsequent flashes advance the remaining manganese ions to higher oxidation states, with an accompanying deprotonation of two bound water molecules. The O–O bond is formed, and oxygen is evolved during the transition from S₃ to S₀ (Fig. 1A). Despite decades of study, many aspects of the water-oxidation mechanism remain to be elucidated. In this work, we obtain previously unknown information concerning proton-coupled electron transfer reactions during the S₁-to-S₂ and other S-state transitions.Open in a separate windowFig. 1.Photosynthetic water oxidation and protonation of a water cluster during the S₁-to-S₂ transition. (A) S-state cycle of photosynthetic water oxidation (7). (A, Inset) Predicted hydrogen-bond network of water molecules in the OEC of PSII (5). Amino acids are shown as sticks. Oxygen atoms of water molecules are shown in blue, with hydrogen bonds to peptide carbonyl groups shown as dashed lines. (B) Mechanism proposed for deprotonation of a terminal water ligand during the S₁-to-S₂ transition in PSII (14). Hydrogen bonds are shown as dashed lines. Putative substrate water molecules are shown in blue. (C) Schematic diagram showing the formation of a cationic water cluster, W₅⁺, on the S₁ to S₂ transition in PSII. (D) Diagram showing method generating the reaction-induced FTIR spectrum, corresponding to the S₂-minus-S₁ spectrum. Difference FTIR spectra for the other S-state transitions are produced with two (S₃-minus-S₂), three (S₀-minus-S₃), or four (S₁–S₀) flashes (see cycle in A).Many mechanisms have been proposed for photosynthetic oxygen evolution (reviewed in refs. 9–13). Fig. 1B shows a possible mechanism for the S₁-to-S₂ transition based on quantum mechanics (QM)/molecular mechanics (MM) calculations (ref. 14; but also see ref. 13). The calcium ion in the metal cluster has been proposed to bind a substrate water molecule and to activate the substrate (15–18). Deprotonation of terminal water ligands is important in decreasing the midpoint potential necessary for oxygen evolution, which is mediated by YZOH/YZ• (midpoint potential; 1V vs. normal hydrogen electrode) and which, therefore, occurs with a low driving force (19).A hydrogen-bonding network containing bound water molecules (Fig. 1 A, Inset) has been assigned in a recent X-ray structure (5). This network has been hypothesized to play a role in the water-oxidizing cycle (18, 20). Recent reaction-induced FTIR studies of the S₁-to-S₂ transition showed that the frequencies of hydrogen-bonded amide C=O groups were markers of hydrogen-bonding changes in the network. In another approach, the recombination kinetics of YZ• were used as a probe of electrostatic changes in the network in the S₂ and S₀ states (21, 22). In both studies, ammonia, a substrate-based inhibitor (23–25), was used to perturb hydrogen bonding in the OEC and was shown to have significant effects on the spectroscopic signals (18, 20, 22).Proton-coupled electron-transfer reactions occur during the S-state cycle (11, 12). Although the S₁-to-S₂ transition is not accompanied by a net proton release to sucrose-containing buffers, proton release accompanies the other S-state transitions (26). Proton-transfer pathways have been proposed based on site-directed mutagenesis and the 3D arrangement of amino acid side chains (5, 27, 28). However, the idea that the water network itself may act as a proton acceptor has not yet been critically evaluated. Spectroscopic signals from protonated water clusters (Fig. 1C) have been identified in model compounds (29) and in proteins (30–33). The OH frequencies of these clusters are red-shifted from bulk water, with a frequency related to the size of the cluster (34, 35). For example, in bacteriorhodopsin, a cluster of internal water molecules acts as a proton donor during the L-to-M transition (36, 37).Here, spectroscopic evidence for the formation of a cationic water cluster, termed W₅⁺, during the S₁-to-S₂ transition is presented (Fig. 1C). The results suggest that deprotonation of a terminal water ligand or a μ–OH bridge occurs on this transition, that the W₅ water cluster acts as a proton acceptor, and that this proton is not released to bulk solvent until later S states. The formation of the protonated cluster is shown to be dependent on temperature, S state, and calcium, consistent with a role for an internal water cluster in photosynthetic oxygen evolution.     

From the Cover: Mode of action uncovered for the specific reduction of methane emissions from ruminants by the small molecule 3-nitrooxypropanol

Ruminants, such as cows, sheep, and goats, predominantly ferment in their rumen plant material to acetate, propionate, butyrate, CO₂, and methane. Whereas the short fatty acids are absorbed and metabolized by the animals, the greenhouse gas methane escapes via eructation and breathing of the animals into the atmosphere. Along with the methane, up to 12% of the gross energy content of the feedstock is lost. Therefore, our recent report has raised interest in 3-nitrooxypropanol (3-NOP), which when added to the feed of ruminants in milligram amounts persistently reduces enteric methane emissions from livestock without apparent negative side effects [Hristov AN, et al. (2015) Proc Natl Acad Sci USA 112(34):10663–10668]. We now show with the aid of in silico, in vitro, and in vivo experiments that 3-NOP specifically targets methyl-coenzyme M reductase (MCR). The nickel enzyme, which is only active when its Ni ion is in the +1 oxidation state, catalyzes the methane-forming step in the rumen fermentation. Molecular docking suggested that 3-NOP preferably binds into the active site of MCR in a pose that places its reducible nitrate group in electron transfer distance to Ni(I). With purified MCR, we found that 3-NOP indeed inactivates MCR at micromolar concentrations by oxidation of its active site Ni(I). Concomitantly, the nitrate ester is reduced to nitrite, which also inactivates MCR at micromolar concentrations by oxidation of Ni(I). Using pure cultures, 3-NOP is demonstrated to inhibit growth of methanogenic archaea at concentrations that do not affect the growth of nonmethanogenic bacteria in the rumen.Since the agricultural and industrial revolution 200 y ago, the methane concentration in the atmosphere has increased from less than 0.6 to 1.8 ppm. The present concentration is only 0.45% of that of CO₂, but because methane has a 28- to 34-fold higher global warming potential than CO₂ on a 100-y horizon, it contributes significantly to global warming (1). On the other hand, the lifetime of atmospheric methane is relatively short relative to CO₂. Accordingly, the climate response to reductions of methane emissions will be relatively rapid. Thus, measures targeting methane emissions are considered paramount to mitigate climate change (2).One of the main anthropogenic sources of atmospheric methane are ruminants (cattle, sheep, goats), the number of which has grown in parallel with the world population. Presently, there are about 1.5 billion cattle, 1.1 billion sheep, and 0.9 billion goats raised by humans (3). Ruminants emit about 100 million tons of methane per year, which corresponds to ∼20% of global methane emissions (4).In the rumen (Fig. 1), plant material is fermented by anaerobic bacteria, protozoa, fungi, and methanogenic archaea in a trophic chain, predominantly yielding acetate, propionate, butyrate, CO₂, and methane with H₂ as intermediate (5, 6). Whereas organic acids are absorbed and metabolized by the animals, methane escapes the rumen into the atmosphere via eructation and breathing of the animals. The generation of methane by methanogenic archaea in the intestine of domestic ruminants lessens feed efficacy, as up to 12% of the gross energy ingested by the animal is lost this way (7).Fig. 1.Methane formation in the rumen of a dairy cow and its inhibition by 3-nitrooxypropanol (3-NOP). The H₂ concentration in the rumen fluid is near 1 µM (≙140 Pa = 0.14% in the gas phase).Methane (CH₄) formation is the main H₂ sink in the rumen. It is formed by methanogenic archaea at the bottom of the trophic chain mainly from carbon dioxide (CO₂) and hydrogen (H₂) (Fig. 1). However, the methane eructated by ruminants contains only minute amounts of H₂; the concentration of dissolved H₂ in the rumen is near 1 µM (8), equivalent to a H₂ partial pressure of near 140 Pa. Because at 1 µM, H₂ formation from most substrates in the rumen is exergonic (9), the low H₂ concentration indicates that H₂ is consumed in the rumen by the methanogens more rapidly than it is formed by other microorganisms (10). The H₂ concentration increases substantially only when methane formation from H₂ and CO₂ is specifically inhibited by more than 50% (10, 11). Already a small increase in the H₂ concentration (8) leads to both down-regulation of H₂-generating pathways (12) and up-regulation of H₂-neutral and H₂-consuming pathways such as propionate formation, resulting in additional energy supply to the host animal (13–15). Thus, the H₂ concentration stays constant, although its consumption by methanogens is partially inhibited in the rumen.The amount of methane formation per unit of ingested feedstuff can differ significantly between individual animals as it is a heritable trait (16). Understanding these differences has been the scientific motivation to pursue the development of selective inhibitors of methanogenesis that are nontoxic to animals (17, 18). Only recently, a compound has been described that apparently can both substantially decrease CH₄ and increase propionate productions in the rumen without compromising animal performance and health (19). It is the small molecule 3-nitrooxypropanol (3-NOP) (chemical structure shown in Fig. 1) that has been found to persistently decrease enteric methane emissions from sheep (20), dairy cows (21), and beef cattle (22) without apparent negative side effects (19). 3-NOP, given to high-producing dairy cows at 60 mg/kg feed dry matter (Fig. 1), not only decreased methane emissions by 30% but also increased body weight gain significantly without negatively affecting feed intake nor milk production and composition (19).Methane formation in methanogenic archaea is catalyzed by methyl-coenzyme M reductase (MCR), involving methyl-coenzyme M and coenzyme B as substrates (Fig. 2A). MCR is a nickel enzyme in which the nickel is bound in a tetrapyrrole derivative named cofactor F₄₃₀ (23, 24). This nickel-containing cofactor has to be in the Ni(I) oxidation state for the enzyme to be active. Because the redox potential E^o′ of the F₄₃₀(Ni²⁺)/ F₄₃₀(Ni¹⁺) couple is −600 mV, the enzyme is very susceptible to inactivation by oxidants (23, 24). MCR has been well characterized by high-resolution X-ray structures (25–27) and EPR spectroscopy (28) with either substrates or products bound.Fig. 2.Binding of 3-NOP to methyl-coenzyme M reductase (MCR) as suggested by molecular docking. The crystal structure of inactive isoenzyme I from M. marburgensis was used in the docking experiments (25). (A) MCR-catalyzed reaction. CH₃-S-CoM, methyl-coenzyme ...The molecular shape of 3-NOP (Fig. 1) is similar to that of methyl-coenzyme M (Fig. 2A). This fact and the moderate oxidation potential of 3-NOP suggested that inhibition of methanogenesis in ruminants is achieved by targeting the active site of MCR, for which we now provide experimental evidence. We start by describing how the development of 3-NOP was facilitated by molecular modeling.     

Quantum delocalization of protons in the hydrogen-bond network of an enzyme active site

Enzymes use protein architectures to create highly specialized structural motifs that can greatly enhance the rates of complex chemical transformations. Here, we use experiments, combined with ab initio simulations that exactly include nuclear quantum effects, to show that a triad of strongly hydrogen-bonded tyrosine residues within the active site of the enzyme ketosteroid isomerase (KSI) facilitates quantum proton delocalization. This delocalization dramatically stabilizes the deprotonation of an active-site tyrosine residue, resulting in a very large isotope effect on its acidity. When an intermediate analog is docked, it is incorporated into the hydrogen-bond network, giving rise to extended quantum proton delocalization in the active site. These results shed light on the role of nuclear quantum effects in the hydrogen-bond network that stabilizes the reactive intermediate of KSI, and the behavior of protons in biological systems containing strong hydrogen bonds.Although many biological processes can be well-described with classical mechanics, there has been much interest and debate as to the role of quantum effects in biological systems ranging from photosynthetic energy transfer, to photoinduced isomerization in the vision cycle and avian magnetoreception (1). For example, nuclear quantum effects, such as tunneling and zero-point energy (ZPE), have been observed to lead to kinetic isotope effects of greater than 100 in biological proton and proton-coupled electron transfer processes (2, 3). However, the role of nuclear quantum effects in determining the ground-state thermodynamic properties of biological systems, which manifest as equilibrium isotope effects, has gained significantly less attention (4).Ketosteroid isomerase (KSI) possesses one of the highest enzyme unimolecular rate constants and thus, is considered a paradigm of proton transfer catalysis in enzymology (5–11). The remarkable rate of KSI is intimately connected to the formation of a hydrogen-bond network in its active site (Fig. 1A), which acts to stabilize a charged dienolate intermediate, lowering its free energy by ∼11 kcal/mol (1 kcal = 4.18 kJ) relative to solution (Fig. S1) (6). This extended hydrogen-bond network in the active site links the substrate to Asp103 and Tyr16, with the latter further hydrogen-bonded to Tyr57 and Tyr32, which is shown in Fig. 1A.Open in a separate windowFig. 1.KSI⋅intermediate and KSI^D40N ? inhibitor complex. Schematic depiction of (A) the KSI⋅intermediate complex during the catalytic cycle (Fig. S1) and (B) a complex between KSI^D40N and phenol, an inhibitor that acts as an intermediate analog. Both the intermediate and the inhibitor are stabilized by a hydrogen-bond network in the active site of KSI. (C) Image of KSI^D40N with the tyrosine triad enlarged and the atoms O16, H16, O32, H32, and O57 labeled (shown with Tyr57 deprotonated) (16).The mutant KSI^D40N preserves the structure of the wild-type (WT) enzyme while mimicking the protonation state of residue 40 in the intermediate complex (Fig. 1B), therefore permitting experimental investigation of an intermediate-like state of the enzyme (6, 12–14). Experiments have identified that, in the absence of an inhibitor, one of the residues in the active site of KSI^D40N is deprotonated (12). Although one might expect the carboxylic acid of Asp103 to be deprotonated, the combination of recent ¹³C NMR and ultraviolet visible spectroscopy (UV-Vis) experiments has shown that the ionization resides primarily on the hydroxyl group of Tyr57, which possesses an anomalously low pK_a of 6.3 ± 0.1 (12). Such a large tyrosine acidity is often associated with specific stabilizing electrostatic interactions (such as a metal ion or cationic residue in close proximity), which is not the case here, suggesting that an additional stabilization mechanism is at play (15).One possible explanation is suggested by the close proximity of the oxygen (O) atoms on the side chains of the adjacent residues Tyr16 (O16) and Tyr32 (O32) to the deprotonated O on Tyr57 (O57) (Fig. 1C) (16). In several high-resolution crystal structures, these distances are found to be around 2.6 Å (14, 16, 17), which is much shorter than those observed in hydrogen-bonded liquids such as water, where O–O distances are typically around 2.85 Å. Such short heavy-atom distances are only slightly larger than those typically associated with low-barrier hydrogen bonds (18–20), where extensive proton sharing is expected to occur between the atoms. In addition, at these short distances, the proton’s position uncertainty (de Broglie wavelength) becomes comparable with the O–O distance, indicating that nuclear quantum effects could play an important role in stabilizing the deprotonated residue (Fig. 1C). In this work, we show how nuclear quantum effects determine the properties of protons in the active-site hydrogen-bond network of KSI^D40N in the absence and presence of an intermediate analog by combining ab initio path integral simulations and isotope effect experiments.     

Direct conversion of plant biomass to ethanol by engineered Caldicellulosiruptor bescii

Ethanol is the most widely used renewable transportation biofuel in the United States, with the production of 13.3 billion gallons in 2012 [John UM (2013) Contribution of the Ethanol Industry to the Economy of the United States]. Despite considerable effort to produce fuels from lignocellulosic biomass, chemical pretreatment and the addition of saccharolytic enzymes before microbial bioconversion remain economic barriers to industrial deployment [Lynd LR, et al. (2008) Nat Biotechnol 26(2):169–172]. We began with the thermophilic, anaerobic, cellulolytic bacterium Caldicellulosiruptor bescii, which efficiently uses unpretreated biomass, and engineered it to produce ethanol. Here we report the direct conversion of switchgrass, a nonfood, renewable feedstock, to ethanol without conventional pretreatment of the biomass. This process was accomplished by deletion of lactate dehydrogenase and heterologous expression of a Clostridium thermocellum bifunctional acetaldehyde/alcohol dehydrogenase. Whereas wild-type C. bescii lacks the ability to make ethanol, 70% of the fermentation products in the engineered strain were ethanol [12.8 mM ethanol directly from 2% (wt/vol) switchgrass, a real-world substrate] with decreased production of acetate by 38% compared with wild-type. Direct conversion of biomass to ethanol represents a new paradigm for consolidated bioprocessing, offering the potential for carbon neutral, cost-effective, sustainable fuel production.Increasing demand for fuels, geopolitical instability, the limitation of global petroleum reserves, and the impact on climate change induced by greenhouse gases have increased the need for renewable and sustainable biofuels (1–5). First-generation biofuels produced from food crops, such as corn, are limited by cost and competition with food supply (6, 7). Switchgrass is a perennial grass native to North America, and its high productivity on marginal farmlands and low agricultural input requirements make it an attractive feedstock for the production of biofuels and biochemicals (8). A yield of 36.7 Mg⋅ha⁻¹ was achieved in field trials in Oklahoma (9), and switchgrass has the potential to produce 500% or more energy than is used for its cultivation (10). The use of abundant lignocellulosic plant biomass as feedstock is environmentally desirable and economically essential for enabling a viable biofuels industry (11). Current strategies for bioethanol production from lignocellulosic feedstocks require three major operational steps: physicochemical pretreatment, enzymatic saccharification, and fermentation (Fig. 1) (6, 12). Pretreatment and enzymatic hydrolysis represent substantial cost and it is estimated that the use of cellulolytic microbes for consolidated bioprocessing and eliminating pretreatment would reduce bioprocessing costs by 40% (2). Considerable effort has been made to develop single microbes capable of both saccharification and fermentation to avoid the substantial expense of using saccharolytic enzyme mixtures (13). Heterologous expression of saccharolytic enzymes has been demonstrated in a number of organisms, including Saccharomyces cerevisiae, Zymomonas mobilis, Escherichia coli, and Bacillus subtilis to ferment various model cellulosic and hemicellulosic substrates (13–15). Although these approaches have resulted in progress in cellulose utilization, the overall enzyme activity is still very low compared with that of naturally cellulolytic organisms and the rates of hydrolysis are not sufficient for an industrial process (13).Open in a separate windowFig. 1.Comparison of bioethanol production strategies and a predicted fermentative pathway in C. bescii. Depiction of “single step bioprocessing” accomplished by engineered C. bescii. CBP, consolidated bioprocessing.High-temperature fermentations facilitate biomass deconstruction and may reduce contamination and volatilize toxic products, such as alcohols. Clostridium thermocellum and Thermoanaerobacterium saccharolyticum have been used in mixed culture fermentations successfully for laboratory-scale demonstration of first-generation consolidated bioprocessing (13, 16) (Fig. 1). C. thermocellum is one promising candidate for consolidated bioprocessing because it is naturally cellulolytic, able to hydrolyze cellulose at 2.5 g⋅L⁻¹⋅h⁻¹, and produces ethanol as one fermentation product, but it has not yet been engineered to produce ethanol at high yield and lacks the ability to ferment hemicellulosic sugars (13, 17). Caldicellulosiruptor bescii, on the other hand, is the most thermophilic cellulolytic bacterium so far described, growing optimally at ∼80 °C with the ability to use a wide range of substrates, such as cellulose, hemicellulose, and lignocellulosic plant biomass without harsh and expensive chemical pretreatment (17, 18), efficiently fermenting both C₅ and C₆ sugars derived from plant biomass (17, 18). C. bescii uses the Embden–Meyerhof–Parnas pathway for conversion of glucose to pyruvate, and the predominant end-products are acetate, lactate, and hydrogen (Fig. 2) (18). A mutant strain of C. bescii (JWCB018) was recently isolated in which the lactate dehydrogenase gene (ldh) was disrupted spontaneously via insertion of a native transposon (19, 20). A complete deletion of ldh was also engineered (21), and this strain no longer produced lactate, instead diverting metabolic flux to additional acetate and H₂, demonstrating the utility of the newly developed tools to provide a platform for further strain engineering. The recent development of genetic methods for the manipulation of this organism (19, 21, 22) opens the door for metabolic engineering for the direct conversion of unpretreated plant biomass to liquid fuels, such as ethanol, via “single step bioprocessing” (Fig. 1).Open in a separate windowFig. 2.Overview of C. bescii fermentative pathways for bioconversion of hexose sugars. Pathway 1 (blue) results in 2 mol of acetic acid and 4 mol of H₂ per mole of glucose. Pathway 2 (green) produces 2 mol of lactic acid per mole of glucose. Pathway 3 (red) is a new pathway resulting from heterologous expression of the C. thermocellum adhE gene to synthesize 2 mol of ethanol per mole of glucose.     

Gate-controlled proton diffusion and protonation-induced ratchet motion in the stator of the bacterial flagellar motor

The proton permeation process of the stator complex MotA/B in the flagellar motor of Escherichia coli was investigated. The atomic model structure of the transmembrane part of MotA/B was constructed based on the previously published disulfide cross-linking and tryptophan scanning mutations. The dynamic permeation of hydronium/sodium ions and water molecule through the channel formed in MotA/B was observed using a steered molecular dynamics simulation. During the simulation, Leu46 of MotB acts as the gate for hydronium ion permeation, which induced the formation of water wire that may mediate the proton transfer to Asp32 on MotB. Free energy profiles for permeation were calculated by umbrella sampling. The free energy barrier for H₃O⁺ permeation was consistent with the proton transfer rate deduced from the flagellar rotational speed and number of protons per rotation, which suggests that the gating is the rate-limiting step. Structure and dynamics of the MotA/B with nonprotonated and protonated Asp32, Val43Met, and Val43Leu mutants in MotB were investigated using molecular dynamics simulation. A narrowing of the channel was observed in the mutants, which is consistent with the size-dependent ion selectivity. In MotA/B with the nonprotonated Asp32, the A3 segment in MotA maintained a kink whereas the protonation induced a straighter shape. Assuming that the cytoplasmic domain not included in the atomic model moves as a rigid body, the protonation/deprotonation of Asp32 is inferred to induce a ratchet motion of the cytoplasmic domain, which may be correlated to the motion of the flagellar rotor.Bacterial flagella are multifuel engines that convert ion motive force to molecular motor rotation. Escherichia coli has a few proton-driven flagellar motors with stators (protein MotA/B complex) in the inner membrane that act as proton channels (1–5). In addition, Vibrio alginolyticus has a polar flagellum powered by sodium ions (6). Bacillus alcalophilus has motors driven by rubidium (Rb⁺), potassium (K⁺), and sodium ions (Na⁺) that can be converted to Na⁺-driven motors by a single mutation (7).The proton transfer mechanism in membrane proteins is associated with water wire and/or a hydrogen bond chain (HBC) (8, 9). The water wire comprises water molecules aligned in a protein channel, where protons are transferred by hopping along the wire. Protons are conducted through the hydrogen bonds formed by the polar amino acid residues and water molecules along the proton transfer pathway in the HBC. Protons can also be transferred by diffusion of hydronium ions (H₃O⁺). The diffusion distance in a hydrophilic environment is short in a liquid (the lifetime in water is ca. 1 ps) (10, 11), but it should be longer in a more hydrophobic environment. H₃O⁺ forms a hydrogen bond (H bond) with the nearest neighbor water molecules and the carbonyl groups, and proton hopping along the H bonds is faster than diffusion of Na⁺ and K⁺ in the ion channel of Gramicidin A (12, 13).These flagellar motors can rotate in both clockwise (CW) and counterclockwise (CCW) directions (viewed from the outside of the cell), and the swimming pattern of the bacteria is controlled by reversal of the motor rotation (14). In E. coli, the FliG, FliM, FliN, MotA, and MotB proteins are involved with torque generation (Fig. 1A) (14, 15). FliG, FliM, and FliN constitute the flagellar rotor and are also involved with the CW/CCW switching. Each rotor is typically surrounded by 10 stators that consist of two membrane proteins, MotA and MotB (PomA and PomB in V. alginolyticus). Each stator is composed of four MotA and two MotB proteins, and can independently produce torque for flagellar rotation.Open in a separate windowFig. 1.Overall structure of MotA/B. (A) Schematic views of the flagellar motors of E. coli (Left) and V. alginolyticus (Right), (B) TM regions modeled, and (C) TM helix arrangement in the initial modeling of MotA/B in E. coli. The obtained atomic model structure viewed parallel to the membrane (D) and from the periplasmic side (E). Spheres denote P atoms in the lipid head groups. MotA/B cross sections of the area enclosed by magenta in E around a channel at the levels of Leu46 (F) and Asp32 (G). x, y, and z axes are defined as in D and E.Systematic Cys and Trp mutagenesis (16–20) has provided essential information on the structure and function of the flagellar motor. Each MotA (295 residues) contains four transmembrane (TM) alpha helical segments (A1–A4), two short loops in the periplasm, and two long segments (residues 61–160 and 228–295) in the cytoplasm (Fig. 1B) (3, 21). Arg90 and Glu98 on the MotA cytoplasmic domain interact with the polar residues on the rotor protein, FliG, during the rotation of the motor (22, 23). It has been suggested that Pro173 and Pro222 at the cytoplasmic sides of A3 and A4 regulate the conformational changes required for torque generation (24). MotB (308 residues) is composed of a short N-terminal cytoplasimic segment, one TM helix (B), and a large C-terminal periplasmic domain (Fig. 1B) (4, 5). Asp32, which is situated near the cytoplasmic end of the B segment, is conserved across the species and considered to be the most plausible proton binding site (25). The B segment is expected to form a proton channel together with A3 and A4 (Fig. 1C) (17). Only a few polar residues have been identified in the predicted TM segments of MotA/B (19, 20), which implies that the channel surface should be relatively hydrophobic. The periplasmic region of MotB has a peptidoglycan binding motif, which anchors the stator complex to the peptidoglycan layer around the rotor (5, 26). Deletion of residues 52–65 just after the B segment causes proton leakage and cell growth arrest, which suggests that this fragment acts as a plug to suppress proton leakage (27). The generation of torque is hypothesized to originate from the conformational changes of the MotA cytoplasmic domain upon proton association/dissociation at the carboxyl group of Asp32 on MotB and by the interaction with FliG in the rotor (28–31).In the present study, the mechanism for proton permeation in MotA/B was investigated (Fig. S1). The atomic structure of MotA/B was constructed based on the disulfide cross-linking (16–18) and tryptophan scanning mutations (19, 20). The dynamic permeation of hydronium ions, sodium ions, and water molecules was observed using a steered molecular dynamics (SMD) simulation (32–34), and free energy profiles for ion/water permeation were calculated by umbrella sampling. The effects of amino acid substitutions related to ion selectivity was investigated, and the possible ratchet motion of the cytoplasmic domain induced by protonation/deprotonation cycle of Asp32 was examined.Open in a separate windowFig. S1.Overall scheme of this study.     

Antimicrobial lipopeptide tridecaptin A1 selectively binds to Gram-negative lipid II

Tridecaptin A₁ (TriA₁) is a nonribosomal lipopeptide with selective antimicrobial activity against Gram-negative bacteria. Here we show that TriA₁ exerts its bactericidal effect by binding to the bacterial cell-wall precursor lipid II on the inner membrane, disrupting the proton motive force. Biochemical and biophysical assays show that binding to the Gram-negative variant of lipid II is required for membrane disruption and that only the proton gradient is dispersed. The NMR solution structure of TriA₁ in dodecylphosphocholine micelles with lipid II has been determined, and molecular modeling was used to provide a structural model of the TriA₁–lipid II complex. These results suggest that TriA₁ kills Gram-negative bacteria by a mechanism of action using a lipid-II–binding motif.Recently, a lot of media coverage has been focused on the problem of antimicrobial resistance. A report commissioned by the UK government predicts that by 2050 antimicrobial resistance will have caused 300 million premature deaths and cost the global economy over $100 trillion (1). Even more worrying is the lack of new classes of antibiotics active against Gram-negative bacteria. In the past 50 y, only a few structurally and mechanistically distinct classes of antibiotics have been clinically approved to treat systemic infections (including fidaxomicin, bedaquiline, linezolid, and daptomycin), yet none of these are active against Gram-negative bacteria (2, 3). Two new classes of Gram-negative targeting antibiotics in the clinical pipeline are POL7080 and brilacidin (4, 5). Both of these compounds are modeled on antimicrobial peptides, which are becoming increasingly important in the fight against antibiotic resistance (6). Bacteria produce a wealth of antimicrobial peptides, both ribosomally, including the lantibiotics (7, 8), and nonribosomally, including lipopeptides (9). In particular, lipopeptides are a rich source of antimicrobial compounds, and several examples with activity against Gram-positive (10, 11) and/or Gram-negative bacteria (12) have been recently characterized.Tridecaptin A₁ (TriA₁) is a member of the tridecaptin family, a group of nonribosomal lipopeptides produced by Bacillus and Paenibacillus species (Fig. 1) (13–15). This acylated tridecapeptide displays strong and selective antimicrobial activity against Gram-negative bacteria, including multidrug-resistant strains of Klebsiella pneumoniae, Acinetobacter baumannii, and Escherichia coli (16). TriA₁ analogs have low cytotoxicity and have been shown to treat K. pneumoniae infections in mice (16, 17). Therefore, we believe that tridecaptin A₁ could be an excellent antibiotic candidate. However, before our investigations little was known about how TriA₁ exerts its selective bactericidal effect against Gram-negative bacteria. A previous structure–activity relationship study by our group suggested that TriA₁, akin to many other lipopeptides, is a membrane-targeting agent. We found that removal of the N-terminal lipid tail abolishes antimicrobial activity; however, the chiral lipid tail could be replaced with an octanoyl chain to give Oct-TriA₁ (Fig. 1), which retains full activity (16). We therefore sought to identify the precise mode and mechanism of action by which TriA₁ kills Gram-negative bacteria.Open in a separate windowFig. 1.Structures of the tridecaptin analogs TriA₁ and Oct-TriA₁.     

Insect nicotinic receptor interactions in vivo with neonicotinoid,organophosphorus, and methylcarbamate insecticides and a synergist

The nicotinic acetylcholine (ACh) receptor (nAChR) is the principal insecticide target. Nearly half of the insecticides by number and world market value are neonicotinoids acting as nAChR agonists or organophosphorus (OP) and methylcarbamate (MC) acetylcholinesterase (AChE) inhibitors. There was no previous evidence for in vivo interactions of the nAChR agonists and AChE inhibitors. The nitromethyleneimidazole (NMI) analog of imidacloprid, a highly potent neonicotinoid, was used here as a radioligand, uniquely allowing for direct measurements of house fly (Musca domestica) head nAChR in vivo interactions with various nicotinic agents. Nine neonicotinoids inhibited house fly brain nAChR [³H]NMI binding in vivo, corresponding to their in vitro potency and the poisoning signs or toxicity they produced in intrathoracically treated house flies. Interestingly, nine topically applied OP or MC insecticides or analogs also gave similar results relative to in vivo nAChR binding inhibition and toxicity, but now also correlating with in vivo brain AChE inhibition, indicating that ACh is the ultimate OP- or MC-induced nAChR active agent. These findings on [³H]NMI binding in house fly brain membranes validate the nAChR in vivo target for the neonicotinoids, OPs and MCs. As an exception, the remarkably potent OP neonicotinoid synergist, O-propyl O-(2-propynyl) phenylphosphonate, inhibited nAChR in vivo without the corresponding AChE inhibition, possibly via a reactive ketene metabolite reacting with a critical nucleophile in the cytochrome P450 active site and the nAChR NMI binding site.The nicotinic nervous system has two principal sites of insecticide action, the nicotinic receptor (nAChR) activated by acetylcholine (ACh) and neonicotinoid agonists (1–6), and acetylcholinesterase (AChE) inhibited by organophosphorus (OP) and methylcarbamate (MC) compounds to generate and maintain localized toxic ACh levels (Fig. 1) (7). The nAChR and AChE targets have been identified in insects by multiple techniques but not by direct assays of the ACh binding site in the brain of poisoned insects. Here we use the outstanding insecticidal potency of the nitromethyleneimidazole (NMI) analog of imidacloprid (IMI) (8) as a radioligand (9), designated [³H]NMI, to directly measure the house fly (Musca domestica) nAChR not only in vitro but also in vivo, allowing us to validate by a previously undescribed method the neonicotinoid direct and OP/MC indirect nAChR targets (Fig. 2). This approach also helped solve the intriguing mechanism by which an O-(2-propynyl) phosphorus compound strongly synergizes neonicotinoid insecticidal activity (10) by dual inhibition of cytochrome P450 (CYP) (11–13) and the nAChR agonist site (described herein). Insecticide disruption at the insect nAChR can now be readily studied in vitro and in vivo with a single radioligand allowing better understanding of the action of several principal insecticide chemotypes (Fig. 3).Open in a separate windowFig. 1.The insect nicotinic receptor is the direct or indirect target for neonicotinoids, organophosphorus compounds and methylcarbamates, which make up about 45% of the insecticides by number and world market value (2, 7).Open in a separate windowFig. 2.In this study, Musca nicotinic receptor in vivo interactions with major insecticide chemotypes are revealed by a [³H]NMI radioligand reporter assay. *Position of tritium label.Open in a separate windowFig. 3.Two neonicotinoid nicotinic agonists and two anticholinesterase insecticides.     

Roles of multiple-proton transfer pathways and proton-coupled electron transfer in the reactivity of the bis-FeIV state of MauG

The high-valent state of the diheme enzyme MauG exhibits charge–resonance (CR) stabilization in which the major species is a bis-Fe^IV state with one heme present as Fe^IV=O and the other as Fe^IV with axial heme ligands provided by His and Tyr side chains. In the absence of its substrate, the high-valent state is relatively stable and returns to the diferric state over several minutes. It is shown that this process occurs in two phases. The first phase is redistribution of the resonance species that support the CR. The second phase is the loss of CR and reduction to the diferric state. Thermodynamic analysis revealed that the rates of the two phases exhibited different temperature dependencies and activation energies of 8.9 and 19.6 kcal/mol. The two phases exhibited kinetic solvent isotope effects of 2.5 and 2.3. Proton inventory plots of each reaction phase exhibited extreme curvature that could not be fit to models for one- or multiple-proton transfers in the transition state. Each did fit well to a model for two alternative pathways for proton transfer, each involving multiple protons. In each case the experimentally determined fractionation factors were consistent with one of the pathways involving tunneling. The percent of the reaction that involved the tunneling pathway differed for the two reaction phases. Using the crystal structure of MauG it was possible to propose proton–transfer pathways consistent with the experimental data using water molecules and amino acid side chains in the distal pocket of the high-spin heme.MauG (1) is a diheme enzyme that catalyzes a six-electron oxidation required for posttranslational modification of a precursor of methylamine dehydrogenase (preMADH) (2) to complete the biosynthesis of its protein-derived cofactor (3) tryptophan tryptophylquinone (TTQ) (4). The hemes of MauG are unusual in several respects. One is a high-spin five-coordinate heme that is ligated by His35. The other is a low-spin six-coordinate heme with two ligands provided by His205 and Tyr294 (1, 5). The latter is, to our knowledge, the first example of natural His–Tyr ligation of a protein-bound heme cofactor, and the first example of Tyr ligation of a c-type heme. An intervening residue, Trp93, “connects” the two hemes (Fig. 1) via rapid electron transfer (ET) (6–9). A unique feature of MauG is that the oxidation of diferric MauG by H₂O₂, or of diferrous MauG by O₂, generates a high-valent bis-Fe^IV state (8) in which the high-spin heme is present as Fe^IV=O with the His35 ligand, and the other heme is present as Fe^IV with the His–Tyr axial ligation retained (5, 10, 11). Formation of the bis-Fe^IV state is accompanied by changes in the visible absorbance spectrum. One observes a decrease in intensity and shift of the Soret peak from 406 to 408 nm and appearance of minor peaks at 526 and 559 nm (Fig. 2) (9, 12).Open in a separate windowFig. 1.Diheme site of MauG. A portion of the crystal structure of the MauG-preMADH complex [Protein Data Bank (PDB) ID code 3L4M] is shown with MauG in pink, the MADH β-subunit in green, and the α subunit in blue. Shown in sticks are the hemes of MauG, the intervening Trp93, the three Met residues that are susceptible to autooxidation, the residues on preMADH that are modified by MauG, and Trp-199 which mediates ET from preMADH to bis-Fe^IV MauG. This figure was produced using PyMOL (www.pymol.org).Open in a separate windowFig. 2.Changes in the absorption spectrum of MauG caused by addition of H₂O₂ to diferric MauG. Spectra of MauG were recorded before (solid line) and after (dashed line) the addition of a stoichiometric amount of H₂O₂.The entire absorbance spectrum (A) is presented and the changes in the Soret region (B) and NIR region (C) are magnified.Despite being a highly potent oxidant, the bis-Fe^IV species displays extraordinary stability with a half-life of several minutes in the absence of its substrate (8). A basis for this stability was inferred from the observation of a near-infrared (NIR) electronic absorption feature centered at 950 nm that was observed in bis-Fe^IV MauG (Fig. 2C). This spectral feature is characteristic of a charge–resonance (CR) transition phenomenon (6, 9). A model was presented in which the CR occurs in the absence of direct heme–heme contact by ultrafast and reversible ET between the two high-valent hemes, via hopping through the intervening Trp93 residue (9). In this model the high-valent form of MauG comprises an ensemble of resonance structures including compound ES-like and compound I-like forms of the hemes, with the bis-Fe^IV as the dominant species.The catalytic mechanism of MauG is unusual in that the preMADH substrate does not make direct contact with either heme but instead binds to the surface of MauG several angstroms away (5). Catalysis requires long-range ET to bis-Fe^IV MauG from the residues on preMADH that are modified via a hole-hopping mechanism through Trp199 (13, 14), which resides at the MauG–preMADH interface (Fig. 1). Concomitant with this ET is the formation of free-radical intermediates on preMADH that go on to form the TTQ product (15). In the absence of preMADH, the autoreduction of the bis-Fe^IV redox state to the diferric state leads to inactivation of MauG (16). Analysis of the damaged MauG revealed that this process involves the oxidation of three Met residues (108, 114, and 116) which are located 7.5–15.2 Å from the high-spin heme iron (Fig. 1) (17).To further investigate the dynamic nature of the ensemble of resonance forms of MauG that comprise the high-valent state and the basis for its stability, temperature-dependence and kinetic solvent isotope effect (KSIE) studies were performed. These studies provide evidence for a redistribution within the ensemble of resonance structures before loss of CR stabilization of the high-valent redox state which is linked to the reduction to the diferric state. Thermodynamic analysis of the rates of reaction of these processes reveals that the rates of the initial redistribution of the ensemble of resonance structures and the subsequent loss of CR stabilization exhibit different dependencies on temperature. This accounts for the fact that the early phase is only observable at lower temperatures. Proton inventories of the KSIE indicate that the rates of both the initial redistribution of the ensemble of high-valent species and the loss of CR stabilization are rate-limited by multiple proton-transfer (PT) steps involving two alternative pathways. The likely pathways are identified from the crystal structure of MauG.     

Taxodione and arenarone inhibit farnesyl diphosphate synthase by binding to the isopentenyl diphosphate site

We used in silico methods to screen a library of 1,013 compounds for possible binding to the allosteric site in farnesyl diphosphate synthase (FPPS). Two of the 50 predicted hits had activity against either human FPPS (HsFPPS) or Trypanosoma brucei FPPS (TbFPPS), the most active being the quinone methide celastrol (IC₅₀ versus TbFPPS ∼20 µM). Two rounds of similarity searching and activity testing then resulted in three leads that were active against HsFPPS with IC₅₀ values in the range of ∼1–3 µM (as compared with ∼0.5 µM for the bisphosphonate inhibitor, zoledronate). The three leads were the quinone methides taxodone and taxodione and the quinone arenarone, compounds with known antibacterial and/or antitumor activity. We then obtained X-ray crystal structures of HsFPPS with taxodione+zoledronate, arenarone+zoledronate, and taxodione alone. In the zoledronate-containing structures, taxodione and arenarone bound solely to the homoallylic (isopentenyl diphosphate, IPP) site, not to the allosteric site, whereas zoledronate bound via Mg²⁺ to the same site as seen in other bisphosphonate-containing structures. In the taxodione-alone structure, one taxodione bound to the same site as seen in the taxodione+zoledronate structure, but the second located to a more surface-exposed site. In differential scanning calorimetry experiments, taxodione and arenarone broadened the native-to-unfolded thermal transition (T_m), quite different to the large increases in ΔT_m seen with biphosphonate inhibitors. The results identify new classes of FPPS inhibitors, diterpenoids and sesquiterpenoids, that bind to the IPP site and may be of interest as anticancer and antiinfective drug leads.Farnesyl diphosphate synthase (FPPS) catalyzes the condensation of isopentenyl diphosphate (IPP; compound 1 in Fig. 1) with dimethylallyl diphosphate (DMAPP; compound 2 in Fig. 1) to form the C₁₀ isoprenoid geranyl diphosphate (GPP; compound 3 in Fig. 1), which then condenses with a second IPP to form the C₁₅ isoprenoid, farnesyl diphosphate (FPP; compound 4 in Fig. 1). FPP then is used in a wide range of reactions including the formation of geranylgeranyl diphosphate (GGPP) (1), squalene (involved in cholesterol and ergosterol biosynthesis), dehydrosqualene (used in formation of the Staphylococcus aureus virulence factor staphyloxanthin) (2), undecaprenyl diphosphate (used in bacterial cell wall biosynthesis), and quinone and in heme a/o biosynthesis. FPP and GGPP also are used in protein (e.g., Ras, Rho, Rac) prenylation, and FPPS is an important target for the bisphosphonate class of drugs (used to treat bone resorption diseases) such as zoledronate (compound 5 in Fig. 1) (3). Bisphosphonates targeting FPPS have activity as antiparasitics (4), act as immunomodulators (activating γδ T cells containing the Vγ2Vδ2 T-cell receptor) (5), and switch macrophages from an M2 (tumor-promoting) to an M1 (tumor-killing) phenotype (6). They also kill tumor cells (7) and inhibit angiogenesis (8). However, the bisphosphonates in clinical use (zoledronate, alendronate, risedronate, ibandronate, etidronate, and clodronate) are very hydrophilic and bind avidly to bone mineral (9). Therefore, there is interest in developing less hydrophilic species (10) that might have better activity against tumors in soft tissues and better antibacterial (11) and antiparasitic activity.Open in a separate windowFig. 1.Chemical structures of FPPS substrates, products, and inhibitors.The structure of FPPS (from chickens) was first reported by Tarshis et al. (12) and revealed a highly α-helical fold. The structures of bacterial and Homo sapiens FPPS (HsFPPS) are very similar; HsFPPS structure (13, 14) is shown in Fig. 2A. There are two substrate-binding sites, called here “S1” and “S2.” S1 is the allylic (DMAPP, GPP) binding site to which bisphosphonates such as zoledronate bind via a [Mg²⁺]₃ cluster (15) (Fig. 2B). S2 is the homoallylic site to which IPP binds, Fig. 2B. Recently, Jahnke et al. (10) and Salcius et al. (16) discovered a third ligand-binding site called the “allosteric site” (hereafter the “A site”). A representative zoledronate+A-site inhibitor structure [Protein Data Bank (PDB) ID code 3N46] (Nov_980; compound 6 in Fig. 1) showing zoledronate in S1 and Nov_980 (compound 6) in the A site is shown in a stereo close-up view in Fig. 2B, superimposed on a zoledronate+IPP structure (PDB ID code 2F8Z) in S2. Whether the allosteric site serves a biological function (e.g., in feedback regulation) has not been reported. Nevertheless, highly potent inhibitors (IC₅₀ ∼80 nM) have been developed (10), and the best of these newly developed inhibitors are far more hydrophobic than are typical bisphosphonates (∼2.4–3.3 for cLogP vs. ∼−3.3 for zoledronate) and are expected to have better direct antitumor effects in soft tissues (10).Open in a separate windowFig. 2.Structures of human FPPS. (A) Structure of HsFPPS showing zoledronate (compound 5) and IPP (compound 1) bound to the S1 (allylic) and S2 (homoallylic) ligand-binding sites (PDB ID code 2F8Z). (B) Superposition of the IPP-zoledronate structure (PDB ID code 2F8Z) on the zoledronate-Nov_980 A-site inhibitor structure (PDB ID code 3N46). Zoledronate binds to the allylic site S1, IPP binds to the homoallylic site S2, and the allosteric site inhibitor binds to the A site. Active-site “DDXXD” residues are indicated, as are Mg²⁺ molecules (green and yellow spheres, respectively). The views are in stereo.In our group we also have developed more lipophilic compounds (e.g., compound 7 in Fig. 1) (17, 18) as antiparasitic (19) and anticancer drug leads (18) and, using computational methods, have discovered other novel nonbisphosphonate FPPS inhibitors (e.g., compound 8 in Fig. 1) that have micromolar activity against FPPS (20). In this study, we extended our computational work and tried to discover other FPPS inhibitors that target the A site. Such compounds would be of interest because they might potentiate the effects of zoledronate and other bisphosphonates, as reported for other FPPS inhibitors (21), and have better tissue distribution properties in general.     

Mutant cycle analysis with modified saxitoxins reveals specific interactions critical to attaining high-affinity inhibition of hNaV1.7

Improper function of voltage-gated sodium channels (Na_Vs), obligatory membrane proteins for bioelectrical signaling, has been linked to a number of human pathologies. Small-molecule agents that target Na_Vs hold considerable promise for treatment of chronic disease. Absent a comprehensive understanding of channel structure, the challenge of designing selective agents to modulate the activity of Na_V subtypes is formidable. We have endeavored to gain insight into the 3D architecture of the outer vestibule of Na_V through a systematic structure–activity relationship (SAR) study involving the bis-guanidinium toxin saxitoxin (STX), modified saxitoxins, and protein mutagenesis. Mutant cycle analysis has led to the identification of an acetylated variant of STX with unprecedented, low-nanomolar affinity for human Na_V1.7 (hNa_V1.7), a channel subtype that has been implicated in pain perception. A revised toxin-receptor binding model is presented, which is consistent with the large body of SAR data that we have obtained. This new model is expected to facilitate subsequent efforts to design isoform-selective Na_V inhibitors.Modulation of action potentials in electrically excitable cells is controlled by tight regulation of ion channel expression and distribution. Voltage-gated sodium ion channels (Na_Vs) constitute one such family of essential membrane proteins, encoded in 10 unique genes (Na_V1.1–Na_V1.9, Na_x) and further processed through RNA splicing, editing, and posttranslational modification. Sodium channels are comprised of a large (∼260 kDa) pore-forming α-subunit coexpressed with ancillary β-subunits. Misregulation and/or mutation of Na_Vs have been ascribed to a number of human diseases including neuropathic pain, epilepsy, and cardiac arrhythmias. A desire to understand the role of individual Na_V subtypes in normal and aberrant signaling motivates the development of small-molecule probes for regulating the function of specific channel isoforms (1–4).Nature has provided a collection of small-molecule toxins, including (+)-saxitoxin (STX, 1) and (−)-tetrodotoxin (TTX), which bind to a subset of mammalian Na_V isoforms with nanomolar affinity (5–7). Guanidinium toxins inhibit Na⁺ influx through Na_Vs by occluding the outer pore above the ion selectivity filter (site 1). This proposed mechanism for toxin block follows from a large body of electrophysiological and site-directed mutagenesis studies (Fig. 1A and refs. 8–10). The detailed view of toxin binding, however, is unsupported by structural biology, as no high-resolution structure of a eukaryotic Na_V has been solved to date (11–16). Na_V homology models, constructed based on X-ray analyses of prokaryotic Na⁺ and K⁺ voltage-gated channels, do not sufficiently account for experimental structure–activity relationship (SAR) data (6, 17–20), and the molecular details underlying distinct differences in toxin potencies toward individual Na_V subtypes remain undefined (5, 6, 21–23). The lack of structural information motivates a comprehensive, systematic study of toxin–protein interactions.Open in a separate windowFig. 1.(A) Schematic drawing of 1 bound in the Na_V outer pore as suggested by previous electrophysiology and mutagenesis experiments. Each of the four domains (I, orange; II, red; III, gray; and IV, teal) is represented by a separate panel. (B) Schematic representation of double-mutant cycle analysis and mathematical definition of coupling energy (ΔΔE_Ω). X¹ = IC₅₀(WT⋅STX)/IC₅₀(MutNa_V⋅STX), X² = IC₅₀(WT⋅MeSTX)/IC₅₀(MutNa_V⋅MeSTX), Y¹ = IC₅₀(MutNa_V⋅STX)/IC₅₀(MutNa_V⋅MeSTX), and Y² = IC₅₀(WT⋅STX)/IC₅₀(WT⋅MeSTX).Double-mutant cycle analysis has proven an invaluable experimental method for assessing protein–protein, protein–peptide, and protein–small-molecule interactions in the absence of crystallographic data (Fig. 1B and Fig. S1 and refs. 9, 10, and 24–31). Herein, we describe mutant cycle analysis with Na_Vs using STX and synthetically modified forms thereof. Our results are suggestive of a toxin–Na_V binding pose distinct from previously published views. Our studies have resulted in the identification of a natural variant of STX that is potent against the STX-resistant human Na_V1.7 isoform (hNa_V1.7). Structural insights gained from these studies provide a foundation for engineering guanidinium toxins with Na_V isoform selectivity.Open in a separate windowFig. S1.Mutant cycle analysis definition and examples. (A) Schematic of a single mutant cycle with mathematical expressions for coupling energy ΔΔE_Ω. R is the ideal gas constant and T is temperature. Each IC₅₀ is the half maximal inhibition concentration determined by whole-cell voltage-clamp electrophysiology. When the separation between IC₅₀ values for the reference compound and the modified compound is different with a mutant than with the WT protein, a nonzero value for ΔΔE_Ω is obtained (B), but when the separation is the same (C), ΔΔE_Ω is equal to 0. In B, the difference in the relative affinity of 1 and 4 with Y401A is smaller than the difference with the WT channel, indicating a positive coupling (ΔΔE_Ω > 0). In C, the relative affinities of 1 and 8 against WT rNa_V1.4 and Y401A are similar, and ΔΔE_Ω ∼0 kcal/mol.