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.     

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.     

Detection of genomic variations and DNA polymorphisms and impact on analysis of meiotic recombination and genetic mapping

DNA polymorphisms are important markers in genetic analyses and are increasingly detected by using genome resequencing. However, the presence of repetitive sequences and structural variants can lead to false positives in the identification of polymorphic alleles. Here, we describe an analysis strategy that minimizes false positives in allelic detection and present analyses of recently published resequencing data from Arabidopsis meiotic products and individual humans. Our analysis enables the accurate detection of sequencing errors, small insertions and deletions (indels), and structural variants, including large reciprocal indels and copy number variants, from comparisons between the resequenced and reference genomes. We offer an alternative interpretation of the sequencing data of meiotic products, including the number and type of recombination events, to illustrate the potential for mistakes in single-nucleotide polymorphism calling. Using these examples, we propose that the detection of DNA polymorphisms using resequencing data needs to account for nonallelic homologous sequences.DNA polymorphisms are ubiquitous genetic variations among individuals and include single nucleotide polymorphisms (SNPs), insertions and deletions (indels), and other larger rearrangements (1–3) (Fig. 1 A and B). They can have phenotypic consequences and also serve as molecular markers for genetic analyses, facilitating linkage and association studies of genetic diseases, and other traits in humans (4–6), animals, plants, (7–10) and other organisms. Using DNA polymorphisms for modern genetic applications requires low-error, high-throughput analytical strategies. Here, we illustrate the use of short-read next-generation sequencing (NGS) data to detect DNA polymorphisms in the context of whole-genome analysis of meiotic products.Open in a separate windowFig. 1.(A) SNPs and small indels between two ecotype genomes. (B) Possible types of SVs. Col genotypes are marked in blue and Ler in red. Arrows indicate DNA segments involved in SVs between the two ecotypes. (C) Meiotic recombination events including a CO and a GC (NCO). Centromeres are denoted by yellow dots.There are many methods for detecting SNPs (11–14) and structural variants (SVs) (15–25), including NGS, which can capture nearly all DNA polymorphisms (26–28). This approach has been widely used to analyze markers in crop species such as rice (29), genes associated with diseases (6, 26), and meiotic recombination in yeast and plants (30, 31). However, accurate identification of DNA polymorphisms can be challenging, in part because short-read sequencing data have limited information for inferring chromosomal context.Genomes usually contain repetitive sequences that can differ in copy number between individuals (26–28, 31); therefore, resequencing analyses must account for chromosomal context to avoid mistaking highly similar paralogous sequences for polymorphisms. Here, we use recently published datasets to describe several DNA sequence features that can be mistaken as allelic (32, 33) and describe a strategy for differentiating between repetitive sequences and polymorphic alleles. We illustrate the effectiveness of these analyses by examining the reported polymorphisms from the published datasets.Meiotic recombination is initiated by DNA double-strand breaks (DSBs) catalyzed by the topoisomerase-like SPORULATION 11 (SPO11). DSBs are repaired as either crossovers (COs) between chromosomes (Fig. 1C), or noncrossovers (NCOs). Both COs and NCOs can be accompanied by gene conversion (GC) events, which are the nonreciprocal transfer of sequence information due to the repair of heteroduplex DNA during meiotic recombination. Understanding the control of frequency and distribution of CO and NCO (including GC) events has important implications for human health (including cancer and aneuploidy), crop breeding, and the potential for use in genome engineering. COs can be detected relatively easily by using polymorphic markers in the flanking sequences, but NCO products can only be detected if they are accompanied by a GC event. Because GCs associated with NCO result in allelic changes at polymorphic sites without exchange of flanking sequences, they are more difficult to detect. Recent advances in DNA sequencing have made the analysis of meiotic NCOs more feasible (30–32, 34); however, SVs present a challenge in these analyses. We recommend a set of guidelines for detection of DNA polymorphisms by using genomic resequencing short-read datasets. These measures improve the accuracy of a wide range of analyses by using genomic resequencing, including estimation of COs, NCOs, and GCs.     

Self-assembly of amphiphilic Janus dendrimers into uniform onion-like dendrimersomes with predictable size and number of bilayers

A constitutional isomeric library synthesized by a modular approach has been used to discover six amphiphilic Janus dendrimer primary structures, which self-assemble into uniform onion-like vesicles with predictable dimensions and number of internal bilayers. These vesicles, denoted onion-like dendrimersomes, are assembled by simple injection of a solution of Janus dendrimer in a water-miscible solvent into water or buffer. These dendrimersomes provide mimics of double-bilayer and multibilayer biological membranes with dimensions and number of bilayers predicted by the Janus compound concentration in water. The simple injection method of preparation is accessible without any special equipment, generating uniform vesicles, and thus provides a promising tool for fundamental studies as well as technological applications in nanomedicine and other fields.Most living organisms contain single-bilayer membranes composed of lipids, glycolipids, cholesterol, transmembrane proteins, and glycoproteins (1). Gram-negative bacteria (2, 3) and the cell nucleus (4), however, exhibit a strikingly special envelope that consists of a concentric double-bilayer membrane. More complex membranes are also encountered in cells and their various organelles, such as multivesicular structures of eukaryotic cells (5) and endosomes (6), and multibilayer structures of endoplasmic reticulum (7, 8), myelin (9, 10), and multilamellar bodies (11, 12). This diversity of biological membranes inspired corresponding biological mimics. Liposomes (Fig. 1) self-assembled from phospholipids are the first mimics of single-bilayer biological membranes (13–16), but they are polydisperse, unstable, and permeable (14). Stealth liposomes coassembled from phospholipids, cholesterol, and phospholipids conjugated with poly(ethylene glycol) exhibit improved stability, permeability, and mechanical properties (17–20). Polymersomes (21–24) assembled from amphiphilic block copolymers exhibit better mechanical properties and permeability, but are not always biocompatible and are polydisperse. Dendrimersomes (25–28) self-assembled from amphiphilic Janus dendrimers and minidendrimers (26–28) have also been elaborated to mimic single-bilayer biological membranes. Amphiphilic Janus dendrimers take advantage of multivalency both in their hydrophobic and hydrophilic parts (23, 29–32). Dendrimersomes are assembled by simple injection (33) of a solution of an amphiphilic Janus dendrimer (26) in a water-soluble solvent into water or buffer and produce uniform (34), impermeable, and stable vesicles with excellent mechanical properties. In addition, their size and properties can be predicted by their primary structure (27). Amphiphilic Janus glycodendrimers self-assemble into glycodendrimersomes that mimic the glycan ligands of biological membranes (35). They have been demonstrated to be bioactive toward biomedically relevant bacterial, plant, and human lectins, and could have numerous applications in nanomedicine (20).Open in a separate windowFig. 1.Strategies for the preparation of single-bilayer vesicles and multibilayer onion-like vesicles.More complex and functional cell mimics such as multivesicular vesicles (36, 37) and multibilayer onion-like vesicles (38–40) have also been discovered. Multivesicular vesicles compartmentalize a larger vesicle (37) whereas multibilayer onion-like vesicles consist of concentric alternating bilayers (40). Currently multibilayer vesicles are obtained by very complex and time-consuming methods that do not control their size (39) and size distribution (40) in a precise way. Here we report the discovery of “single–single” (28) amphiphilic Janus dendrimer primary structures that self-assemble into uniform multibilayer onion-like dendrimersomes (Fig. 1) with predictable size and number of bilayers by simple injection of their solution into water or buffer.     

De novo production of the plant-derived alkaloid strictosidine in yeast

The monoterpene indole alkaloids are a large group of plant-derived specialized metabolites, many of which have valuable pharmaceutical or biological activity. There are ∼3,000 monoterpene indole alkaloids produced by thousands of plant species in numerous families. The diverse chemical structures found in this metabolite class originate from strictosidine, which is the last common biosynthetic intermediate for all monoterpene indole alkaloid enzymatic pathways. Reconstitution of biosynthetic pathways in a heterologous host is a promising strategy for rapid and inexpensive production of complex molecules that are found in plants. Here, we demonstrate how strictosidine can be produced de novo in a Saccharomyces cerevisiae host from 14 known monoterpene indole alkaloid pathway genes, along with an additional seven genes and three gene deletions that enhance secondary metabolism. This system provides an important resource for developing the production of more complex plant-derived alkaloids, engineering of nonnatural derivatives, identification of bottlenecks in monoterpene indole alkaloid biosynthesis, and discovery of new pathway genes in a convenient yeast host.Monoterpene indole alkaloids (MIAs) are a diverse family of complex nitrogen-containing plant-derived metabolites (1, 2). This metabolite class is found in thousands of plant species from the Apocynaceae, Loganiaceae, Rubiaceae, Icacinaceae, Nyssaceae, and Alangiaceae plant families (2, 3). Many MIAs and MIA derivatives have medicinal properties; for example, vinblastine, vincristine, and vinflunine are approved anticancer therapeutics (4, 5). These structurally complex compounds can be difficult to chemically synthesize (6, 7). Consequently, industrial production relies on extraction from the plant, but these compounds are often produced in small quantities as complex mixtures, making isolation challenging, laborious, and expensive (8–10). Reconstitution of plant pathways in microbial hosts is proving to be a promising approach to access plant-derived compounds as evidenced by the successful production of terpenes, flavonoids, and benzylisoquinoline alkaloids in microorganisms (11–19). Microbial hosts can also be used to construct hybrid biosynthetic pathways to generate modified natural products with potentially enhanced bioactivities (8, 20, 21). Across numerous plant species, strictosidine is believed to be the core scaffold from which all 3,000 known MIAs are derived (1, 2). Strictosidine undergoes a variety of redox reactions and rearrangements to form the thousands of compounds that comprise the MIA natural product family (Fig. 1) (1, 2). Due to the importance of strictosidine, the last common biosynthetic intermediate for all known MIAs, we chose to focus on heterologous production of this complex molecule (1). Therefore, strictosidine reconstitution represents the necessary first step for heterologous production of high-value MIAs.Open in a separate windowFig. 1.Strictosidine, the central intermediate in monoterpene indole alkaloid (MIA) biosynthesis, undergoes a series of reactions to produce over 3,000 known MIAs such as vincristine, quinine, and strychnine.     

A family of starch-active polysaccharide monooxygenases

The recently discovered fungal and bacterial polysaccharide monooxygenases (PMOs) are capable of oxidatively cleaving chitin, cellulose, and hemicelluloses that contain β(1→4) linkages between glucose or substituted glucose units. They are also known collectively as lytic PMOs, or LPMOs, and individually as AA9 (formerly GH61), AA10 (formerly CBM33), and AA11 enzymes. PMOs share several conserved features, including a monocopper center coordinated by a bidentate N-terminal histidine residue and another histidine ligand. A bioinformatic analysis using these conserved features suggested several potential new PMO families in the fungus Neurospora crassa that are likely to be active on novel substrates. Herein, we report on NCU08746 that contains a C-terminal starch-binding domain and an N-terminal domain of previously unknown function. Biochemical studies showed that NCU08746 requires copper, oxygen, and a source of electrons to oxidize the C1 position of glycosidic bonds in starch substrates, but not in cellulose or chitin. Starch contains α(1→4) and α(1→6) linkages and exhibits higher order structures compared with chitin and cellulose. Cellobiose dehydrogenase, the biological redox partner of cellulose-active PMOs, can serve as the electron donor for NCU08746. NCU08746 contains one copper atom per protein molecule, which is likely coordinated by two histidine ligands as shown by X-ray absorption spectroscopy and sequence analysis. Results indicate that NCU08746 and homologs are starch-active PMOs, supporting the existence of a PMO superfamily with a much broader range of substrates. Starch-active PMOs provide an expanded perspective on studies of starch metabolism and may have potential in the food and starch-based biofuel industries.Polysaccharide monooxygenases (PMOs) are enzymes secreted by a variety of fungal and bacterial species (1–5). They have recently been found to oxidatively degrade chitin (6–8) and cellulose (8–14). PMOs have been shown to oxidize either the C1 or C4 atom of the β(1→4) glycosidic bond on the surface of chitin (6, 7) or cellulose (10–12, 14), resulting in the cleavage of this bond and the creation of new chain ends that can be subsequently processed by hydrolytic chitinases and cellulases. Several fungal PMOs were shown to significantly enhance the degradation of cellulose by hydrolytic cellulases (9), indicating that these enzymes can be used in the conversion of plant biomass into biofuels and other renewable chemicals.There are three families of PMOs characterized thus far: fungal PMOs that oxidize cellulose (9–12) (also known as GH61 and AA9); bacterial PMOs that are active either on chitin (6, 8) or cellulose (8, 13) (also known as CBM33 and AA10); and fungal PMOs that oxidize chitin (AA11) (7). Sequence homology between these three families is very low. Nevertheless, the available structures of PMOs from all three families reveal a conserved fold, including an antiparallel β-sandwich core and a highly conserved monocopper active site on a flat protein surface (Fig. 1A) (2, 6, 7, 9, 10, 15–17). Two histidine residues in a motif termed the histidine brace coordinate the copper center. The N-terminal histidine ligand binds in a bidentate mode, and its imidazole ring is methylated at the N_ε position in fungal PMOs (Fig. 1A).Open in a separate windowFig. 1.(A) Representative overall and active site structures of fungal PMOs (PDB ID code 2YET) (10). (B) Structure of cellulose (18, 19). Chitin also contains β(1→4) linkages and has similar crystalline higher order structure to cellulose. (C) Model structure of amylopectin (23–25). Hydrogen bonds are shown with green dashed lines.Considering the conserved structural features, it is not surprising that the currently known PMOs act on substrates with similar structures. Cellulose and chitin contain long linear chains of β(1→4) linked glucose units and N-acetylglucosamine units, respectively (Fig. 1B). The polymer chains form extensive hydrogen bonding networks, which result in insoluble and very stable crystalline structures (18–21). PMOs are thought to bind to the substrate with their flat active site surface, which orients the copper center for selective oxidation at the C1 or C4 position (6, 16, 22). Some bacterial chitin-binding proteins are cellulose-active PMOs (8, 13, 14), further suggesting that the set of PMO substrates is restricted to β(1→4) linked polymers of glucose and glucose derivatives.Here, we report on the identification of new families of PMOs that contain several key features of previously characterized PMOs, but act on substrates different from cellulose or chitin. A member of one of these novel families of PMOs, NCU08746, was shown to oxidatively cleave amylose, amylopectin, and starch. We designate the NCU08746 family as starch-active PMOs. Both amylose and amylopectin contain linear chains of α(1→4) linked glucose, whereas the latter also contains α(1→6) glycosidic linkages at branch points in the otherwise α(1→4) linked polymer. Unlike cellulose and chitin, amylose and amylopectin do not form microcrystals; instead, they exist in disordered, single helical, and double helical forms (23–27) (see Fig. 1C for example). Starch exists partially in nanocrystalline form, but lacks the flat molecular surfaces as those found in chitin and cellulose. The discovery of starch-active PMOs shows that this oxidative mechanism of glycosidic bond cleavage is more widespread than initially expected.     

Thermodynamic origins of protein folding,allostery, and capsid formation in the human hepatitis B virus core protein

HBc, the capsid-forming “core protein” of human hepatitis B virus (HBV), is a multidomain, α-helical homodimer that aggressively forms human HBV capsids. Structural plasticity has been proposed to be important to the myriad functions HBc mediates during viral replication. Here, we report detailed thermodynamic analyses of the folding of the dimeric HBc protomer under conditions that prevented capsid formation. Central to our success was the use of ion mobility spectrometry–mass spectrometry and microscale thermophoresis, which allowed folding mechanisms to be characterized using just micrograms of protein. HBc folds in a three-state transition with a stable, dimeric, α-helical intermediate. Extensive protein engineering showed thermodynamic linkage between different structural domains. Unusual effects associated with mutating some residues suggest structural strain, arising from frustrated contacts, is present in the native dimer. We found evidence of structural gatekeepers that, when mutated, alleviated native strain and prevented (or significantly attenuated) capsid formation by tuning the population of alternative native conformations. This strain is likely an evolved feature that helps HBc access the different structures associated with its diverse essential functions. The subtle balance between native and strained contacts may provide the means to tune conformational properties of HBc by molecular interactions or mutations, thereby conferring allosteric regulation of structure and function. The ability to trap HBc conformers thermodynamically by mutation, and thereby ablate HBV capsid formation, provides proof of principle for designing antivirals that elicit similar effects.The “protein-folding problem” describes how a polypeptide sequence contains all the information needed for it to adopt a specific 3D structure spontaneously (1). The chemistry and thermodynamic code that causes proteins to fold also underpins protein–protein interactions, allostery, and supramolecular assembly. An emerging trend has been the study of model proteins free from kinetic traps, aggregation, or metal binding, features that can confound experimental execution and data interpretation (2, 3). Consequently, model proteins are small (typically <130 residues), soluble monomers with few proline or cysteine residues and no prosthetic groups (2, 3).Although model proteins have been instrumental in taking the field to its current zenith, there is a paucity of experimental insights into the conformational dynamics of larger, oligomeric proteins, especially those implicated in diseases (3). Such proteins usually have complex behavior refractory to detailed experimental studies. However, the connection between sequence, structure, dynamics, and allostery makes studies of larger proteins central to understanding biological function and aiding drug design (vide infra) (4). One such protein is HBc, the capsid-forming “core protein” of human hepatitis B virus (HBV), a major pathogen that kills 600,000 people annually (5). Although excellent vaccines exist, there are no effective cures for extant chronic infections (5, 6). In addition to capsid formation, HBc plays many essential roles in HBV replication (7–9), making it an attractive drug target (10–15).WT HBc is a 183-residue polypeptide comprising a structured capsid-forming region (residues 1–149; Fig. 1A) and a basic, nucleic acid-binding domain (residues 150–183) (16–18). The structured N-terminal region (hereafter HBc_1–149) spontaneously self-assembles in vitro and in vivo to form icosahedral capsid-like particles (CLPs) identical to nucleocapsids isolated from patient serum (19, 20). X-ray crystallography and cryo-EM have characterized the structure of HBc_1–149 within the context of CLPs, virions, and hexamers (16, 19–23). HBc homodimers comprise two structural domains (Fig. 1A): Helices α₃ and α₄ from opposing monomers pack together and form a disulfide-linked, four-helix bundle dimerization interface (visible as protrusions on the capsid exterior; Fig. 1B), whereas α₁, α₂, and α₅ pack together and around the base of the four-helix bundle to create the hydrophobic core of “contact” domains (19). Weak interdimer interactions between contact domains stabilize HBV capsids (19, 24) (Fig. 1B).Open in a separate windowFig. 1.HBc_1–149 dimer structure within HBV capsids. (A) Four-helix bundle dimerization interface (black) is flanked by contact domains (orange and red). Helices are numbered, and the N and C termini of one monomer are indicated. The disulfide link between C61 of each monomer is indicated (cyan). (B) Exterior surface of a T = 4 capsid HBc_1–149 (PDB ID code 1QGT) (19). Dimers around the threefold and fivefold axes are indicated in blue/green and purple/orange, respectively. (Inset) Interacting quasiequivalent HBc_1–149 dimers from the fivefold (purple and orange) and threefold (blue and green) axes are shown. Hydrophobic contacts between contact domains stabilize capsids. Residues that perturb capsid formation when mutated are indicated.Multiple studies show clearly that HBc has a very malleable structure, with this structural plasticity argued to be functionally important (22, 23). This hypothesis accords well with antivirals that modulate HBc structure (11–15, 22, 23). Studies of HBV capsid assembly have inferred the existence of assembly-active (HBc^Ass) and assembly-incompetent (HBc^Inc) HBc conformations (12, 13, 21, 24, 25). However, there are few detailed insights on the thermodynamic origins of structure, allostery, and dynamics for the dimeric HBc_1–149 protomer, where structural plasticity must originate. This arises from dimeric HBc_1–149 being very challenging to study in vitro (compared with the model proteins described above) because it is a 298-residue disulfide-linked homodimer (containing 6 cysteine and 24 proline residues) that aggregates aggressively and forms capsids.Here, we report detailed folding and stability studies of dimeric HBc_1–149. These show HBc_1–149 folds in a three-state transition with a populated, dimeric, α-helical intermediate. Of 29 “chemically conservative” mutants used to probe folding energetics (26), many had similar effects on the stability of the intermediate and native ensembles. The distribution of these mutations was consistent with the intermediate being stabilized by a significant native-like structure. However, some mutations destabilized the native state (N) much less than the intermediate state (I) relative to the denatured state (D), or significantly increased the free energy of unfolding (ΔG_D–N) relative to WT HBc_1–149. This suggests HBc_1–149 contains structural strain arising from frustrated contacts (27, 28). We found evidence of HBc_1–149 adopting multiple native conformers, where capsid assembly-competent conformers were less stable than those incapable of, or attenuated in, capsid formation. Frustrated regions likely contain structural gatekeepers that (28), when mutated, subtly tuned the folding energy landscape and altered capsid assembly. The presence of multiple native conformations and frustrated regions may explain the origins of allostery reported for HBc. Frustration is likely an evolved tradeoff that balances the conflicting requirements of HBc folding with allosteric regulation of native structure, capsid formation, and diverse functions of different conformers (29). The ability to trap HBc conformers thermodynamically by mutation and ablate capsid formation provides a proof of principle for designing antivirals that elicit similar effects.     

Single-molecule spectroscopy reveals how calmodulin activates NO synthase by controlling its conformational fluctuation dynamics

Mechanisms that regulate the nitric oxide synthase enzymes (NOS) are of interest in biology and medicine. Although NOS catalysis relies on domain motions, and is activated by calmodulin binding, the relationships are unclear. We used single-molecule fluorescence resonance energy transfer (FRET) spectroscopy to elucidate the conformational states distribution and associated conformational fluctuation dynamics of the two electron transfer domains in a FRET dye-labeled neuronal NOS reductase domain, and to understand how calmodulin affects the dynamics to regulate catalysis. We found that calmodulin alters NOS conformational behaviors in several ways: It changes the distance distribution between the NOS domains, shortens the lifetimes of the individual conformational states, and instills conformational discipline by greatly narrowing the distributions of the conformational states and fluctuation rates. This information was specifically obtainable only by single-molecule spectroscopic measurements, and reveals how calmodulin promotes catalysis by shaping the physical and temporal conformational behaviors of NOS.Although proteins adopt structures determined by their amino acid sequences, they are not static objects and fluctuate among ensembles of conformations (1). Transitions between these states can occur on a variety of length scales (Å to nm) and time scales (ps to s) and have been linked to functionally relevant phenomena such as allosteric signaling, enzyme catalysis, and protein–protein interactions (2–4). Indeed, protein conformational fluctuations and dynamics, often associated with static and dynamic inhomogeneity, are thought to play a crucial role in biomolecular functions (5–11). It is difficult to characterize such spatially and temporally inhomogeneous dynamics in bulk solution by an ensemble-averaged measurement, especially in proteins that undergo multiple-conformation transformations. In such cases, single-molecule spectroscopy is a powerful approach to analyze protein conformational states and dynamics under physiological conditions, and can provide a molecular-level perspective on how a protein’s structural dynamics link to its functional mechanisms (12–21).A case in point is the nitric oxide synthase (NOS) enzymes (22–24), whose nitric oxide (NO) biosynthesis involves electron transfer reactions that are associated with relatively large-scale movement (tens of Å) of the enzyme domains (Fig. 1A). During catalysis, NADPH-derived electrons first transfer into an FAD domain and an FMN domain in NOS that together comprise the NOS reductase domain (NOSr), and then transfer from the FMN domain to a heme group that is bound in a separate attached “oxygenase” domain, which then enables NO synthesis to begin (22, 25–27). The electron transfers into and out of the FMN domain are the key steps for catalysis, and they appear to rely on the FMN domain cycling between electron-accepting and electron-donating conformational states (28, 29) (Fig. 1B). In this model, the FMN domain is suggested to be highly dynamic and flexible due to a connecting hinge that allows it to alternate between its electron-accepting (FAD→FMN) or closed conformation and electron-donating (FMN→heme) or open conformation (Fig. 1 A and B) (28, 30–36). In the electron-accepting closed conformation, the FMN domain interacts with the NADPH/FAD domain (FNR domain) to receive electrons, whereas in the electron donating open conformation the FMN domain has moved away to expose the bound FMN cofactor so that it may transfer electrons to a protein acceptor like the NOS oxygenase domain, or to a generic protein acceptor like cytochrome c. In this way, the reductase domain structure cycles between closed and open conformations to deliver electrons, according to a conformational equilibrium that determines the movements and thus the electron flux capacity of the FMN domain (25, 28, 32, 34, 35, 37). A similar conformational switching mechanism is thought to enable electron transfer through the FMN domain in the related flavoproteins NADPH-cytochrome P450 reductase and methionine synthase reductase (38–42).Open in a separate windowFig. 1.(A) The nNOSr ribbon structure (from PDB: 1TLL) showing bound FAD (yellow) in FNR domain (green), FMN (orange) in FMN domain (yellow), connecting hinge (blue), and the Cy3–Cy5 label positions (pink) and distance (42 Å, dashed line). (B) Cartoon of an equilibrium between the FMN-closed and FMN-open states, with Cy dye label positions indicated. (C) Cytochrome c reductase activity of nNOSr proteins in their CaM-bound and CaM-free states. Color scheme of bar graphs: Black, WT nNOSr unlabeled; Red, Cys-lite (CL) nNOSr unlabeled; Blue, E827C/Q1268C CL nNOSr unlabeled; and Dark cyan, E827C/Q1268C CL nNOSr labeled.NOS enzymes also contain a calmodulin (CaM) binding domain that is located just before the N terminus of the FMN domain (Fig. 1B), and this provides an important layer of regulation (25, 27). CaM binding to NOS enzymes increases electron transfer from NADPH through the reductase domain and also triggers electron transfer from the FMN domain to the NOS heme as is required for NO synthesis (31, 32). The ability of CaM, or similar signaling proteins, to regulate electron transfer reactions in enzymes is unusual, and the mechanism is a topic of interest and intensive study. It has long been known that CaM binding alters NOSr structure such that, on average, it populates a more open conformation (43, 44). Recent equilibrium studies have detected a buildup of between two to four discreet conformational populations in NOS enzymes and in related flavoproteins, and in some cases, have also estimated the distances between the bound FAD and FMN cofactors in the different species (26, 36, 37, 39, 40), and furthermore, have confirmed that CaM shifts the NOS population distribution toward more open conformations (34, 36, 45). Although valuable, such ensemble-averaged results about conformational states cannot explain how electrons transfer through these enzymes, or how CaM increases the electron flux in NOS, because answering these questions requires a coordinate understanding of the dynamics of the conformational fluctuations. Indeed, computer modeling has indicated that a shift toward more open conformations as is induced by CaM binding to nNOS should, on its own, actually diminish electron flux through nNOS and through certain related flavoproteins (38). Despite its importance, measuring enzyme conformational fluctuation dynamics is highly challenging, and as far as we know, there have been no direct measures on the NOS enzymes or on related flavoproteins, nor studies on how CaM binding might influence the conformational fluctuation dynamics in NOS.To address this gap, we used single-molecule fluorescence energy resonance transfer (FRET) spectroscopy to characterize individual molecules of nNOSr that had been labeled at two specific positions with Cyanine 3 (Cy3) donor and Cyanine 5 (Cy5) acceptor dye molecules, regarding their conformational states distribution and the associated conformational fluctuation dynamics, which in turn enabled us to determine how CaM binding impacts both parameters. This work provides a unique perspective and a novel study of the NOS enzymes and within the broader flavoprotein family, which includes the mammalian enzymes methionine synthase reductase (MSR) and cytochrome P450 reductase (CPR), and reveals how CaM’s control of the conformational behaviors may regulate the electron transfer reactions of NOS catalysis.     

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.     

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.     

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.