Evidence that Ubiquinone Is a Required Intermediate for Rhodoquinone Biosynthesis in Rhodospirillum rubrum

Rhodoquinone (RQ) is an important cofactor used in the anaerobic energy metabolism of Rhodospirillum rubrum. RQ is structurally similar to ubiquinone (coenzyme Q or Q), a polyprenylated benzoquinone used in the aerobic respiratory chain. RQ is also found in several eukaryotic species that utilize a fumarate reductase pathway for anaerobic respiration, an important example being the parasitic helminths. RQ is not found in humans or other mammals, and therefore inhibition of its biosynthesis may provide a parasite-specific drug target. In this report, we describe several in vivo feeding experiments with R. rubrum used for the identification of RQ biosynthetic intermediates. Cultures of R. rubrum were grown in the presence of synthetic analogs of ubiquinone and the known Q biosynthetic precursors demethylubiquinone, demethoxyubiquinone, and demethyldemethoxyubiquinone, and assays were monitored for the formation of RQ₃. Data from time course experiments and S-adenosyl-l-methionine-dependent O-methyltransferase inhibition studies are discussed. Based on the results presented, we have demonstrated that Q is a required intermediate for the biosynthesis of RQ in R. rubrum.Rhodospirillum rubrum is a well-characterized and metabolically diverse member of the family of purple nonsulfur bacteria (29, 61). R. rubrum is typically found in aquatic environments and can adapt to a variety of growth conditions by using photosynthesis, respiration, or fermentation pathways (28, 70). In the light, R. rubrum exhibits photoheterotrophic growth using organic substrates or photoautotrophic growth using CO₂ and H₂ (15, 70). In the dark, R. rubrum can utilize either aerobic respiration (70, 73) or anaerobic respiration with a fumarate reduction pathway or with nonfermentable substrates in the presence of oxidants such as dimethyl sulfoxide (DMSO) or trimethylamine oxide (15, 58, 73). R. rubrum can also grow anaerobically in the dark by fermentation of sugars in the presence of bicarbonate (58). The focus of this work was the biosynthesis of quinones used by R. rubrum for aerobic and anaerobic respiration.Rhodoquinone (RQ; compound 1 in Fig. ​Fig.1)1) is an aminoquinone structurally similar to ubiquinone (coenzyme Q or Q [compound 2]) (44); however, the two differ considerably in redox potential (that of RQ is −63 mV, and that of Q is +100 mV) (2). Both RQ and Q have a fully substituted benzoquinone ring and a polyisoprenoid side chain that varies in length (depending on the species; see Fig. ​Fig.11 for examples). The only difference between the structures is that RQ has an amino substituent (NH₂) instead of a methoxy substituent (OCH₃) on the quinone ring. While Q is a ubiquitous lipid component involved in aerobic respiratory electron transport (9, 36, 60), RQ functions in anaerobic respiration in R. rubrum (19) and in several other phototrophic purple bacteria (21, 22, 41) and is also present in a few aerobic chemotrophic bacteria, including Brachymonas denitrificans and Zoogloea ramigera (23). In these varied species of bacteria, RQ has been proposed to function in fumarate reduction to maintain NAD⁺/NADH redox balance, either during photosynthetic anaerobic metabolism (12, 15-18, 64) or in chemotrophic metabolism when the availability of oxygen as a terminal oxidant is limiting (23). Another recent finding is that RQH₂ is capable of inducing Q-cycle bypass reactions in the cytochrome bc₁ complex in Saccharomyces cerevisiae, resulting in superoxide formation (7). If RQ/RQH₂ coexists in the cytoplasmic membrane with Q/QH₂ in R. rubrum, it might serve as both a substrate for and an inhibitor of the bc₁ complex (47).Open in a separate windowFIG. 1.Proposed pathways for RQ biosynthesis. The number of isoprene units (n) varies by species (in S. cerevisiae, n = 6; in E. coli, n = 8; in C. elegans, n = 9; in helminth parasites, n = 9 or 10; in R. rubrum, n = 10; in humans, n = 10). RQ is not found in S. cerevisiae, E. coli, or humans. Known Coq (from S. cerevisiae) and Ubi (from E. coli) gene products required for the biosynthesis of ubiquinone (Q, compound 2) are labeled. A polyisoprenyl diphosphate (compound 5) is assembled from dimethylallyl disphosphate (compound 3) and isopentyl diphosphate (compound 4). Coupling of compound 5 with p-hydroxybenzoic acid (compound 6) yields 3-polyprenyl-4-hydroxybenzoic acid (compound 7). The next three steps differ between S. cerevisiae and E. coli. However, they merge at the common intermediate (compound 8), which is oxidized to demethyldemethoxyubiquinone (DDMQ_n, compound 9). RQ (compound 1) has been proposed to arise from compound 9, demethoxyubiquinone (DMQ_n; compound 10), demethylubiquinone (DMeQ_n; compound 11), or compound 2 (by pathway A, B, C, or D). Results presented in this work support pathway D as the favored route for RQ biosynthesis in R. rubrum.RQ is also found in the mitochondrial membrane of eukaryotic species capable of fumarate reduction, such as the flagellate Euglena gracilis (25, 53), the free-living nematode Caenorhabditis elegans (62), and the parasitic helminths (65, 66, 68, 72). Similar to R. rubrum, these species can adapt their metabolism to both aerobic and anaerobic conditions throughout their life cycle. For example, most adult parasitic species (e.g., Ascaris suum, Fasciola hepatica, and Haemonchus contortus) rely heavily on fumarate reduction for their energy generation while inside a host organism, where the oxygen tension is very low (30, 65, 72). Under these conditions, the biosynthesis of RQ is upregulated; however, during free-living stages of their life cycle, the helminth parasites use primarily aerobic respiration, which requires Q (30, 65, 72). The anaerobic energy metabolism of the helminthes has been reviewed (63, 67). Humans and other mammalian hosts use Q for aerobic energy metabolism but do not produce or require RQ; therefore, selective inhibition of RQ biosynthesis may lead to highly specific antihelminthic drugs that do not have a toxic effect on the host (35, 48).R. rubrum is an excellent facultative model system for the study of RQ biosynthesis. The complete genome of R. rubrum has recently been sequenced by the Department of Energy Joint Genome Institute, finished by the Los Alamos Finishing Group, and further validated by optical mapping (57). The 16S rRNA sequence of R. rubrum is highly homologous to cognate eukaryotic mitochondrial sequences (46). Due to the similarities in structure, the biosynthetic pathways of RQ and Q have been proposed to diverge from a common precursor (67). Proposed pathways for RQ biosynthesis (A to D), in conjunction with the known steps in Q biosynthesis, are outlined in Fig. ​Fig.11 (31, 34, 60). Parson and Rudney previously showed that when R. rubrum was grown anaerobically in the light in the presence of [U-¹⁴C]p-hydroxybenzoate, ¹⁴C was incorporated into both Q₁₀ and RQ₁₀ (50). In their growth experiments, the specific activity of Q₁₀ was measured at its maximal value 15 h after inoculation and then began to decrease. However, the specific activity of RQ₁₀ continued to increase for 40 h before declining. These results suggested that Q₁₀ was a biosynthetic precursor of RQ₁₀, although this was not directly demonstrated using radiolabeled Q₁₀; hence, the possibility remained that the labeled RQ₁₀ was derived from another radiolabeled lipid species. We have done this feeding experiment with a synthetic analog of Q where n = 3 (Q₃) and monitored for the production of RQ₃. The synthesis and use of farnesylated quinone and aromatic intermediates for characterization of the Q biosynthetic pathway in S. cerevisiae and Escherichia coli has been well documented (4, 5, 38, 52, 59). The other proposed precursors of RQ shown in Fig. ​Fig.11 were also fed to R. rubrum, and the lipid extracts from these assays were analyzed for the presence of RQ₃, i.e., demethyldemethoxyubiquinone-3 (DDMQ₃; compound 9), demethoxyubiquinone-3 (DMQ₃; compound 10), and demethylubiquinone-3 (DMeQ₃; compound 11).In S. cerevisiae and E. coli, the last O-methylation step in Q biosynthesis is catalyzed by the S-adenosyl-l-methionine (SAM)-dependent methyltransferases Coq3 and UbiG, respectively (26, 52); this final methylation step converts DMeQ to Q. Using the NCBI Basic Local Alignment Search Tool, an O-methyltransferase (GeneID no. 3834724 Rru_A0742) that had 41% and 59% sequence identity with Coq3 and UbiG, respectively, was identified in R. rubrum. S-Adenosyl-l-homocysteine (SAH) is a well-known inhibitor of SAM-dependent methyltransferases (13, 24). Because SAH is the transmethylation by-product of SAM-dependent methyltransferases, it is not readily taken up by cells and must be generated in vivo (24). SAH can be produced in vivo from S-adenosine and l-homocysteine thiolactone by endogenous SAH hydrolase (SAHH) (37, 71). A search of the R. rubrum genome also confirmed the presence of a gene encoding SAHH (GeneID no. 3836896 Rru_A3444). It was proposed that if DMeQ is the immediate precursor of RQ, then SAH inhibition of the methyltransferase required for Q biosynthesis should have little effect on RQ production. Conversely, if Q is required for RQ synthesis, then inhibition of Q biosynthesis should have a significant effect on RQ production. Assays were designed to quantify the levels of RQ₃ produced from DMeQ₃ and Q₃ in R. rubrum cultures at various concentrations of SAH.     

Requirements for Construction of a Functional Hybrid Complex of Photosystem I and [NiFe]-Hydrogenase

The development of cellular systems in which the enzyme hydrogenase is efficiently coupled to the oxygenic photosynthesis apparatus represents an attractive avenue to produce H₂ sustainably from light and water. Here we describe the molecular design of the individual components required for the direct coupling of the O₂-tolerant membrane-bound hydrogenase (MBH) from Ralstonia eutropha H16 to the acceptor site of photosystem I (PS I) from Synechocystis sp. PCC 6803. By genetic engineering, the peripheral subunit PsaE of PS I was fused to the MBH, and the resulting hybrid protein was purified from R. eutropha to apparent homogeneity via two independent affinity chromatographical steps. The catalytically active MBH-PsaE (MBH_PsaE) hybrid protein could be isolated only from the cytoplasmic fraction. This was surprising, since the MBH is a substrate of the twin-arginine translocation system and was expected to reside in the periplasm. We conclude that the attachment of the additional PsaE domain to the small, electron-transferring subunit of the MBH completely abolished the export competence of the protein. Activity measurements revealed that the H₂ production capacity of the purified MBH_PsaE fusion protein was very similar to that of wild-type MBH. In order to analyze the specific interaction of MBH_PsaE with PS I, His-tagged PS I lacking the PsaE subunit was purified via Ni-nitrilotriacetic acid affinity and subsequent hydrophobic interaction chromatography. Formation of PS I-hydrogenase supercomplexes was demonstrated by blue native gel electrophoresis. The results indicate a vital prerequisite for the quantitative analysis of the MBH_PsaE-PS I complex formation and its light-driven H₂ production capacity by means of spectroelectrochemistry.Molecular hydrogen (H₂) is often discussed as an alternative source of energy (13, 22, 26, 41). It is a highly energetic, renewable, and zero-carbon dioxide emission fuel; however, it is produced mainly from fossil resources. One intriguing possibility for sustainable H₂ production is the development of cellular systems in which the light-driven oxygenic photosynthesis is efficiently coupled to hydrogen production by hydrogenase (1, 21, 36).During the process of oxygenic photosynthesis, photosystem II (PS II), a thylakoid membrane (TM)-embedded multiprotein complex, utilizes solar energy to oxidize water into dioxygen (O₂), protons, and electrons. The electrons released by PS II are further conducted through an electron transport chain consisting of plastoquinones, the cytochrome b₆f complex, and either plastocyanin or cytochrome c₆ to the chlorophyll (Chl) dimer P₇₀₀ in photosystem I (PS I) (20, 48). During light-induced charge separation in PS I, P₇₀₀ is oxidized, leading to the reduction of the adjacent cofactor A₀ (Chl a). From there, the electrons are transmitted to the phylloquinone A₁ and subsequently to the Fe₄S₄ clusters F_X, F_A, and F_B (9) that are located at the acceptor site of PS I. The acceptor site is composed of the PsaC subunit, which harbors the iron-sulfur clusters F_A and F_B, and the two additional cofactor-free extrinsic subunits PsaD and PsaE. In the final step, the electrons are transferred from F_B to the ferredoxin (PetF), which has a midpoint potential of −412 mV (see Fig. ​Fig.1B)1B) (8, 9).Open in a separate windowFIG. 1.Models of the hydrogenase and photosystem I complexes used in this study. (A) Membrane-bound hydrogenase (MBH_wt) of Ralstonia eutropha H16. (B) Wild-type photosystem I (PS I) from Synechocystis sp. PCC 6803. (C) MBH_stop protein lacking the C-terminal anchor domain of HoxK. (D) MBH_PsaE and PS I_ΔPsaE.Hydrogenases of the NiFe and FeFe types catalyze the reversible cleavage of H₂ into protons and electrons (18, 63). For most hydrogenases, this reaction is highly sensitive to O₂ and leads to the reversible or even irreversible inactivation of the enzyme (49, 66, 67). A prominent exception is the oxygen-tolerant membrane-bound [NiFe]-hydrogenase (MBH) from Ralstonia eutropha H16, which catalyzes H₂ conversion in the presence of O₂ (42, 65). The MBH consists of large subunit HoxG (67 kDa), harboring the NiFe active site, and small subunit HoxK (35 kDa), bearing three FeS clusters (Fig. ​(Fig.1)1) (32). Both cofactor-containing subunits are completely assembled within the cytoplasm and become subsequently translocated through the cytoplasmic membrane by the twin-arginine translocation (Tat) system. This transport is guided by a specific Tat signal peptide that is located at the N terminus of small subunit HoxK (53). The MBH is then connected to the membrane via the hydrophobic C-terminal “anchor” domain of HoxK, which provides the electronic connection to the diheme cytochrome b, HoxZ (5, 57). All structural, accessory, and regulatory genes for the synthesis of active MBH are arranged in a large, megaplasmid-borne operon (7, 11, 14, 29, 33, 38, 58).The concept of light-driven hydrogen production has been investigated in numerous studies (for reviews, see references 3, 21, and 23), including one involving direct electron transfer from PS I to the free form of hydrogenase in vitro (45). In a preliminary attempt, the MBH from R. eutropha was recently directly fused to PsaE (creating MBH_PsaE) (28). The fusion protein was partially purified and subjected to in vitro reconstitution with PS I lacking PsaE (PS I_ΔPsaE) (54) for light-driven hydrogen production. This concept was based on the previous observation that PS I lacking the peripheral subunit PsaE is fully reconstituted in vitro simply by the addition of independently purified PsaE protein (12).In the present communication, we describe a novel purification procedure for R. eutropha MBH_PsaE that yields homogeneous, functionally active MBH_PsaE. Additionally, a new method for efficient and fast purification of Synechocystis sp. PCC 6803 (hereafter referred to as Synechocystis) His-tagged PS I was established. Finally, the pure proteins MBH_PsaE and PS I_ΔPsaE were successfully subjected to in vitro reconstitution.     

Inhibition of the Activity of Both Domains of Lysostaphin through Peptidoglycan Modification by the Lysostaphin Immunity Protein

Resistance to lysostaphin, a staphylolytic glycylglycine endopeptidase, is due to a FemABX-like immunity protein that inserts serines in place of some glycines in peptidoglycan cross bridges. These modifications inhibit both binding of the recombinant cell wall targeting domain and catalysis by the recombinant catalytic domain of lysostaphin.Lysostaphin is a glycylglycine endopeptidase produced by Staphylococcus simulans biovar staphylolyticus (18) that lyses susceptible staphylococci by hydrolyzing the polyglycine cross bridges in their cell wall peptidoglycans (3). The lysostaphin gene sequence was independently determined in 1987 by two groups (8, 13). BLAST analysis (1) of mature lysostaphin revealed two domains: an N-terminal catalytic domain (CAT), which is a member of the M23 family of zinc metalloendopeptidases, and a C-terminal cell wall targeting domain (CWT), which is a member of the SH3b domain family (Fig. ​(Fig.11 A).Open in a separate windowFIG. 1.(A) Schematic diagram of mature lysostaphin, the recombinant catalytic domain (rCAT) (lysostaphin residues 1 to 148), and the recombinant cell wall targeting domain (rCWT) (lysostaphin residues 149 to 246). The numbers represent the beginning and end of the domains, and the solid boxes indicate the N-terminal His₆ tag of the recombinant proteins. (B) SDS-PAGE analysis of rCAT and rCWT purified by a nickel affinity column. Mobilities of molecular mass standards are given on the left side of the gel.The lysostaphin endopeptidase resistance gene (epr or lif) encodes a FemABX-like immunity protein that is located adjacent to the lysostaphin gene on the plasmid pACK1 in S. simulans bv. staphylolyticus (4, 7, 20). Members of the FemABX family of proteins are nonribosomal peptidyl transferases that are involved in the addition of cross bridge amino acids during peptidoglycan subunit synthesis in the cytoplasm (15). In S. simulans bv. staphylolyticus, the lysostaphin immunity protein inserts serines in place of some glycines during peptidoglycan synthesis, which provides resistance to lysostaphin (4, 20).Originally it was suggested that the incorporation of serines in these peptidoglycan cross bridges gave increased resistance to lysostaphin because of the inability of the enzyme to hydrolyze glycyl-serine or seryl-glycine bonds (4, 14, 16). Others later reported that the CWT specifically binds to the polyglycine cross bridges in staphylococci (6) and the binding of CWT to producer-strain cells was less than that to susceptible cells (2). However, the ability of the enzyme or its targeting domain to bind to purified peptidoglycans from staphylococci containing the lysostaphin resistance gene has not been determined. Therefore, we determined if the modification to staphylococcal peptidoglycan cross bridges made by the lysostaphin immunity protein affected the activity of the binding domain, the catalytic domain, or both.     

The S Helix Mediates Signal Transmission as a HAMP Domain Coiled-Coil Extension in the NarX Nitrate Sensor from Escherichia coli K-12

In the nitrate-responsive, homodimeric NarX sensor, two cytoplasmic membrane α-helices delimit the periplasmic ligand-binding domain. The HAMP domain, a four-helix parallel coiled-coil built from two α-helices (HD1 and HD2), immediately follows the second transmembrane helix. Previous computational studies identified a likely coiled-coil-forming α-helix, the signaling helix (S helix), in a range of signaling proteins, including eucaryal receptor guanylyl cyclases, but its function remains obscure. In NarX, the HAMP HD2 and S-helix regions overlap and apparently form a continuous coiled-coil marked by a heptad repeat stutter discontinuity at the distal boundary of HD2. Similar composite HD2-S-helix elements are present in other sensors, such as Sln1p from Saccharomyces cerevisiae. We constructed deletions and missense substitutions in the NarX S helix. Most caused constitutive signaling phenotypes. However, strongly impaired induction phenotypes were conferred by heptad deletions within the S-helix conserved core and also by deletions that remove the heptad stutter. The latter observation illuminates a key element of the dynamic bundle hypothesis for signaling across the heptad stutter adjacent to the HAMP domain in methyl-accepting chemotaxis proteins (Q. Zhou, P. Ames, and J. S. Parkinson, Mol. Microbiol. 73:801-814, 2009). Sequence comparisons identified other examples of heptad stutters between a HAMP domain and a contiguous coiled-coil-like heptad repeat sequence in conventional sensors, such as CpxA, EnvZ, PhoQ, and QseC; other S-helix-containing sensors, such as BarA and TorS; and the Neurospora crassa Nik-1 (Os-1) sensor that contains a tandem array of alternating HAMP and HAMP-like elements. Therefore, stutter elements may be broadly important for HAMP function.Transmembrane signaling in homodimeric bacterial sensors initiates upon signal ligand binding to the extracytoplasmic domain. In methyl-accepting chemotaxis proteins (MCPs), the resulting conformational change causes a displacement of one transmembrane α-helix (TM α-helix) relative to the other. This motion is conducted by the HAMP domain to control output domain activity (reviewed in references 33 and 39).Certain sensors of two-component regulatory systems share topological organization with MCPs. For example, the paralogous nitrate sensors NarX and NarQ contain an amino-terminal transmembrane signaling module similar to those in MCPs, in which a pair of TM α-helices delimit the periplasmic ligand-binding domain (Fig. ​(Fig.1)1) (24) (reviewed in references 32 and 62). The second TM α-helix connects to the HAMP domain. Hybrid proteins in which the NarX transmembrane signaling module regulates the kinase control modules of the MCPs Tar, DifA, and FrzCD demonstrate that NarX and MCPs share a mechanism for transmembrane signaling (73, 74, 81, 82).Open in a separate windowFIG. 1.NarX modular structure. Linear representation of the NarX protein sequence, from the amino (N) to carboxyl (C) termini, drawn to scale. The four modules are indicated at the top of the figure and shown in bold typeface, whereas domains within each module are labeled with standard (lightface) typeface. The nomenclature for modules follows that devised by Swain and Falke (67) for MCPs. Overlap between the HAMP domain HD2 and S-helix elements is indicated in gray. The three conserved Cys residues within the central module (62) are indicated. TM1 and TM2 denote the two transmembrane helices. Helices H1 to H4 of the periplasmic domain (24), and the transmitter domain H, N, D, G (79), and X (41) boxes, are labeled. The HPK 7 family of transmitter sequences, including NarX, have no F box and an unconventional G box (79). The scale bar at the bottom of the figure shows the number of aminoacyl residues.The HAMP domain functions as a signal conversion module in a variety of homodimeric proteins, including histidine protein kinases, adenylyl cyclases, MCPs, and certain phosphatases (12, 20, 77). This roughly 50-residue domain consists of a pair of amphiphilic α-helices, termed HD1 and HD2 (formerly AS1 and AS2) (67), joined by a connector (Fig. ​(Fig.2A).2A). Results from nuclear magnetic resonance (NMR) and electron paramagnetic resonance (EPR) spectroscopy, Cys and disulfide scanning, and mutational analysis converge on a model in which the HD1 and HD2 α-helices form a four-helix parallel coiled-coil (7, 20, 30, 42, 67, 75, 84). The mechanisms through which HAMP domains mediate signal conduction remain to be established (30, 42, 67, 84) (for commentary, see references 43, 49, and 50).Open in a separate windowFIG. 2.HAMP domain extensions. (A) Sequences from representative MCPs (E. coli Tsr and Salmonella enterica serovar Typhimurium Tar) and S-helix-containing sensors (E. coli NarX, NarQ, and BarA, and S. cerevisiae Sln1p). The HAMP domain, S-helix element, and the initial sequence of the MCP adaptation region are indicated. Flanking numbers denote positions of the terminal residues within the overall sequence. Sequential heptad repeats are indicated in alternating bold and standard (lightface) typeface. Numbering for heptad repeats in the methylation region and S-helix sequences has been described previously (4, 8). Numbers within the HD1 and HD2 helices indicate interactions within the HAMP domain (42). Residues at heptad positions a and d are enclosed within boxes, residues at the stutter position a/d are enclosed within a thickly outlined box, and residues in the S-helix ERT signature are in bold typeface. (B) NarX mutational alterations. Deletions are depicted as boxes, and missense substitutions are shown above the sequence. Many of these deletions were reported previously (10) and are presented here for comparison. The phenotypes conferred by the alterations are indicated as follows: impaired induction, black box; constitutive and elevated basal, light gray box; reversed response, dark gray box; wild-type, white box; null, striped box.Coiled-coils result from packing of two or more α-helices (27). The primary sequence of coiled-coils exhibits a characteristic heptad repeat pattern, denoted as a-b-c-d-e-f-g (52, 61), in which positions a and d are usually occupied by nonpolar residues (reviewed in references 1, 47, and 80). For example, the coiled-coil nature of the HAMP domain can be seen in the heptad repeat patterns within the HD1 and HD2 sequences (Fig. ​(Fig.2A2A).Coiled-coil elements adjacent to the HAMP domain have been identified in several sensors, including Saccharomyces cerevisiae Sln1p (69) and Escherichia coli NarX (60). Recently, this element was defined as a specific type of dimeric parallel coiled-coil, termed the signaling helix (S helix), present in a wide range of signaling proteins (8). Sequence comparisons delimit a roughly 40-residue element with a conserved heptad repeat pattern (Fig. ​(Fig.2A).2A). Based on mutational analyses of Sln1p and other proteins, the S helix is suggested to function as a switch that prevents constitutive activation of adjacent output domains (8).The term “signaling helix” previously was used to define the α4-TM2 extended helix in MCPs (23, 33). Here, we use the term S helix to denote the element described by Anantharaman et al. (8).The NarX and NarQ sensors encompass four distinct modules (Fig. ​(Fig.1):1): the amino-terminal transmembrane signaling module, the signal conversion module (including the HAMP domain and S-helix element), the central module of unknown function, and the carboxyl-terminal transmitter module (62). The S-helix element presumably functions together with the HAMP domain in conducting ligand-responsive motions from the transmembrane signaling module to the central module, ultimately regulating transmitter module activity.Regulatory output by two-component sensors reflects opposing transmitter activities (reviewed in reference 55). Positive function results from transmitter autokinase activity; the resulting phosphosensor serves as a substrate for response regulator autophosphorylation. Negative function results from transmitter phosphatase activity, which accelerates phosphoresponse regulator autodephosphorylation (reviewed in references 64 and 65). We envision a homogeneous two-state model for NarX (17), in which the equilibrium between these mutually exclusive conformations is modulated by ligand-responsive signaling.Previous work from our laboratory concerned the NarX and other HAMP domains (9, 10, 26, 77) and separately identified a conserved sequence in NarX and NarQ sensors, the Y box, that roughly corresponds to the S helix (62). Therefore, we were interested to explore the NarX S helix and to test some of the predictions made for its function. Results show that the S helix is critical for signal conduction and suggest that it functions as an extension of the HAMP HD2 α-helix in a subset of sensors exemplified by Sln1p and NarX. Moreover, a stutter discontinuity in the heptad repeat pattern was found to be essential for the NarX response to signal and to be conserved in several distinct classes of HAMP-containing sensors.     

Tryptophan Residues in the Portal Protein of Herpes Simplex Virus 1 Critical to the Interaction with Scaffold Proteins and Incorporation of the Portal into Capsids

Incorporation of the herpes simplex virus 1 (HSV-1) portal vertex into the capsid requires interaction with a 12-amino-acid hydrophobic domain within capsid scaffold proteins. The goal of this work was to identify domains and residues in the U_L6-encoded portal protein pU_L6 critical to the interaction with scaffold proteins. We show that whereas the wild-type portal and scaffold proteins readily coimmunoprecipitated with one another in the absence of other viral proteins, truncation beyond the first 18 or last 36 amino acids of the portal protein precluded this coimmunoprecipitation. The coimmunoprecipitation was also precluded by mutation of conserved tryptophan (W) residues to alanine (A) at positions 27, 90, 127, 163, 241, 262, 532, and 596 of U_L6. All of these W-to-A mutations precluded the rescue of a viral deletion mutant lacking U_L6, except W163A, which supported replication poorly, and W596A, which fully rescued replication. A recombinant virus bearing the W596A mutation replicated and packaged DNA normally, and scaffold proteins readily coimmunoprecipitated with portal protein from lysates of infected cells. Thus, viral functions compensated for the W596A mutation''s detrimental effects on the portal-scaffold interaction seen during transient expression of portal and scaffold proteins. In contrast, the W27A mutation precluded portal-scaffold interactions in infected cell lysates, reduced the solubility of pU_L6, decreased incorporation of the portal into capsids, and abrogated viral-DNA cleavage and packaging.Immature herpesvirus capsids or procapsids consist of two shells: an inner shell, or scaffold, and an outer shell that is roughly spherical and largely composed of the major capsid protein VP5 (24, 38).The capsid scaffold consists of a mixture of the U_L26.5 and U_L26 gene products, with the U_L26.5 gene product (pU_L26.5, ICP35, or VP22a) being the most abundant (1, 12, 20, 21, 32, 38). The U_L26.5 open reading frame shares its coding frame and C terminus with the U_L26 gene but initiates at codon 307 of U_L26 (17). The extreme C termini of both VP22a and the UL26-encoded protein (pU_L26) interact with the N terminus of VP5 (7, 14, 26, 40, 41). Capsid assembly likely initiates when the portal binds VP5/VP22a and/or VP5/pU_L26 complexes (22, 25). The addition of more of these complexes to growing capsid shells eventually produces a closed sphere bearing a single portal. pU_L26 within the scaffold contains a protease that cleaves itself between amino acids 247 and 248, separating pU_L26 into an N-terminal protease domain called VP24 and a C-terminal domain termed VP21 (4, 5, 8, 9, 28, 42). The protease also cleaves 25 amino acids from pU_L26 and VP22a to release VP5 (5, 8, 9). VP21 and VP22a are replaced with DNA when the DNA is packaged (12, 29).When capsids undergo maturation, the outer protein shell angularizes to become icosahedral (13). One fivefold-symmetrical vertex in the angularized outer capsid shell is biochemically distinct from the other 11 and is called the portal vertex because it serves as the channel through which DNA is inserted as it is packaged (23). In herpes simplex virus (HSV), the portal vertex is composed of 12 copies of the portal protein encoded by U_L6 (2, 23, 39). We and others have shown that interactions between scaffold and portal proteins are critical for incorporation of the portal into the capsid (15, 33, 44, 45). Twelve amino acids of scaffold proteins are sufficient to interact with the portal protein, and tyrosine and proline resides within this domain are critical for the interaction with scaffold proteins and incorporation of the portal into capsids (45).One goal of the current study was to map domains and residues within the U_L6-encoded portal protein that mediate interaction with scaffold proteins. We show that the portal-scaffold interaction requires all but the first 18 and last 36 amino acids of pU_L6, as well as several tryptophan residues positioned throughout the portal protein.     

RNA Structures Required for Production of Subgenomic Flavivirus RNA

Flaviviruses are a group of single-stranded, positive-sense RNA viruses causing ∼100 million infections per year. We have recently shown that flaviviruses produce a unique, small, noncoding RNA (∼0.5 kb) derived from the 3′ untranslated region (UTR) of the genomic RNA (gRNA), which is required for flavivirus-induced cytopathicity and pathogenicity (G. P. Pijlman et al., Cell Host Microbe, 4: 579-591, 2008). This RNA (subgenomic flavivirus RNA [sfRNA]) is a product of incomplete degradation of gRNA presumably by the cellular 5′-3′ exoribonuclease XRN1, which stalls on the rigid secondary structure stem-loop II (SL-II) located at the beginning of the 3′ UTR. Mutations or deletions of various secondary structures in the 3′ UTR resulted in the loss of full-length sfRNA (sfRNA1) and production of smaller and less abundant sfRNAs (sfRNA2 and sfRNA3). Here, we investigated in detail the importance of West Nile virus Kunjin (WNV_KUN) 3′ UTR secondary structures as well as tertiary interactions for sfRNA formation. We show that secondary structures SL-IV and dumbbell 1 (DB1) downstream of SL-II are able to prevent further degradation of gRNA when the SL-II structure is deleted, leading to production of sfRNA2 and sfRNA3, respectively. We also show that a number of pseudoknot (PK) interactions, in particular PK1 stabilizing SL-II and PK3 stabilizing DB1, are required for protection of gRNA from nuclease degradation and production of sfRNA. Our results show that PK interactions play a vital role in the production of nuclease-resistant sfRNA, which is essential for viral cytopathicity in cells and pathogenicity in mice.Arthropod-borne flaviviruses such as West Nile virus (WNV), dengue virus (DENV), and Japanese encephalitis virus (JEV) cause major outbreaks of potentially fatal disease and affect over 50 million people every year. The highly pathogenic North American strain of WNV (WNV_NY99) has already claimed more than 1,000 lives with over 27,000 cases reported since its emergence in New York in 1999 and has raised global public health concerns (9). In contrast, the closely related Australian strain of WNV, WNV_KUN, is highly attenuated and does not cause overt disease in humans and animals (11). WNV_KUN has been used extensively as a model virus to study flavivirus replication and flavivirus-host interactions (13, 14, 16-19, 26, 38, 39).The ∼11-kb positive-stranded, capped WNV genomic RNA (gRNA) lacks a poly(A) tail and consists of 5′ and 3′ untranslated regions (UTRs) flanking one open reading frame, which encodes the viral proteins required for the viral life cycle (6, 15, 38, 39). Flavivirus UTRs are involved in translation and initiation of RNA replication and likely determine genome packaging (13, 14, 16, 21, 30, 39-41). Both the 5′ UTR (∼100 nucleotides [nt] in size) and the 3′ UTR (from ∼400 to 700 nucleotides) can form secondary and tertiary structures which are highly conserved among mosquito-borne flaviviruses (1, 8, 10, 14, 29, 32, 34). More specifically, the WNV_KUN 3′ UTR consists of several conserved regions and secondary structures (Fig. ​(Fig.1A)1A) which were previously predicted or shown to exist in various flaviviruses by computational and chemical analyses, respectively (4, 10, 25, 26, 29-32). The 5′ end of the 3′ UTR starts with an AU-rich region which can form stem-loop structure I (SL-I) followed by SL-II, which we previously showed to be vitally important for subgenomic flavivirus RNA (sfRNA) production (26; see also below). SL-II is followed by a short, repeated conserved hairpin (RCS3) and SL-III (26). Further downstream of SL-III are the SL-IV and CS3 structures, which are remarkably similar to the preceding SL-II-RCS3 structure (26, 29). Further downstream of the SL-IV-CS3 structure are dumbbells 1 and 2 (DB1 and DB2, respectively) followed by a short SL and the 3′ SL (25, 26).Open in a separate windowFIG. 1.(A) Model of the WNV_KUN 3′ UTR RNA structure. Highlighted in bold are the secondary structures investigated here. Dashed lines indicate putative PKs. The two sites of the putative PK interactions are shown in open boxes. sfRNA1, -2, -3, and -4 start sites are indicated by arrows. (R)CS, (repeated) conserved sequence; DB, dumbbell structure; PK, pseudoknot; SL, stem-loop. (B) Structural model of PK1 in SL-II with disruptive mutations. Nucleotide numbering is from the end of the 3′ UTR. The sfRNA1 start is indicated by an arrow. Nucleotides forming PK1 are on a gray background, and mutated nucleotides are white on a black background. (C) Sequences mutated in the different constructs. Nucleotides in the wt PK sequences used for mutations are bold and underlined. Introduced mutations are shown under the corresponding nucleotides in the wt sequence.The described structures have been investigated in some detail for their requirement in RNA replication and translation. Generally, a progressive negative effect on viral growth was shown with progressive deletions into the 3′-proximal region of the JEV 3′ UTR (41). However, only a relatively short region of the JEV 3′ UTR, consisting of the 3′-terminal 193 nt, was shown to be absolutely essential for gRNA replication (41). The minimal region for DENV replication was reported to be even shorter (23). Extensive analysis has shown that the most 3′-terminal, essential regions of the 3′ UTR include the cyclization sequence and 3′ SL, which are required for efficient RNA replication (2, 14, 16, 23, 35). As we showed, deletion of SL-II or SL-I did not overtly affect WNV_KUN replication (26). However, deletion of CS2, RCS2, CS3, or RCS3 in WNV replicon RNA significantly reduced RNA replication but not translation (20), indicating that these elements facilitate but are not essential for RNA replication. In addition, it was shown that deletion of DB1 or DB2 resulted in a viable mutant virus that was reduced in growth efficiency, while deletion of both DB structures resulted in a nonviable mutant (23).In addition to the above-mentioned secondary stem-loop structures, computational and chemical analysis of the flavivirus 3′ UTR suggested the presence of 5 pseudoknot (PK) interactions (Fig. ​(Fig.1A)1A) (25, 26, 32). A PK is a structure formed upon base pairing of a single-stranded region of RNA in the loop of a hairpin to a stretch of complementary nucleotides elsewhere in the RNA chain (Fig. ​(Fig.1B).1B). These structures are referred to as hairpin type (H-type) PKs (3), and they usually stabilize secondary RNA structures. Typically, the final tertiary structure does not significantly alter the preformed secondary structure (5). In general, PK interactions have been shown to be important in biological processes such as initiation and/or elongation of translation, initiation of gRNA replication, and ribosomal frameshifting for a number of different viruses, including flaviviruses (reviewed in references 3 and 22). The first PK in the WNV 3′ UTR was predicted to form in SL-II, followed by a similar PK in SL-IV (26) (PK1 and PK2 in Fig. ​Fig.1A).1A). For the DENV, yellow fever virus (YFV), and JEV subgroup of flaviviruses, two PKs further downstream were predicted to form between DB1 and DB2 and corresponding single-stranded RNA regions located further downstream (25) (PK3 and PK4 in Fig. ​Fig.1A).1A). The formation of these structures is supported by covariations in the WNV RNAs. In addition, a PK was proposed to form between a short SL and the 3′ SL at the 3′ terminus of the viral genome (32) (PK5 in Fig. ​Fig.1A1A).Importantly, in addition to its role in viral replication and translation, we have shown that the WNV_KUN 3′ UTR is important for the production of a small noncoding RNA fragment designated sfRNA (26). This short RNA fragment of ∼0.5 kb is derived from the 3′ UTR of the gRNA and exclusively produced by the members of the Flavivirus genus of the Flaviviridae family, where it is required for efficient viral replication, cytopathicity, and pathogenicity (26). Our studies suggested that sfRNA is a product of incomplete degradation of the gRNA presumably by the cellular 5′-3′ exoribonuclease XRN1, resulting from XRN1 stalling on the rigid secondary/tertiary structures located at the beginning of the 3′ UTR (26). XRN1 is an exoribonuclease which usually degrades mRNA from the 5′ to the 3′ end as part of cellular mRNA decay and turnover (33), and it was shown previously that XRN1 can be stalled by SL structures (28). Mutations or deletions of WNV 3′ UTR secondary structures resulted in the loss of full-length sfRNA (sfRNA1) and production of smaller and less abundant sfRNAs (sfRNA2 and sfRNA3) (26). In particular, SL-II (Fig. ​(Fig.1A)1A) was shown to be important for sfRNA1 production; deletion of this structure either alone or in conjunction with other structures located downstream of SL-II abolished sfRNA1 production, leading to the production of the smaller RNA fragments sfRNA2 and sfRNA3.Here, we extended our investigation and studied the importance of several predicted 3′ UTR secondary structures and PK interactions for the production of sfRNA. To further understand the generation mechanism of sfRNA and its requirements, we deleted or mutated a number of RNA structures in the WNV_KUN 3′ UTR and investigated the size and amount of sfRNA generated from these mutant RNAs. The results show that not only SLs but also PK interactions play a vital role in stabilizing the 3′ UTR RNA and preventing complete degradation of viral gRNA to produce nuclease-resistant sfRNA, which is required for efficient virus replication and cytopathicity in cells and virulence in mice.     

Metabolic Engineering of the Tricarboxylic Acid Cycle for Improved Lysine Production by Corynebacterium glutamicum

In the present work, lysine production by Corynebacterium glutamicum was improved by metabolic engineering of the tricarboxylic acid (TCA) cycle. The 70% decreased activity of isocitrate dehydrogenase, achieved by start codon exchange, resulted in a >40% improved lysine production. By flux analysis, this could be correlated to a flux shift from the TCA cycle toward anaplerotic carboxylation.With an annual market volume of more than 1,000,000 tons, lysine is one of the dominating products in biotechnology. In recent years, rational metabolic engineering has emerged as a powerful tool for lysine production (16, 18, 22). Hereby, different target enzymes and pathways in the central metabolism could be identified and successfully modified to create superior production strains (1, 2, 5, 8, 10, 17-20). The tricarboxylic acid (TCA) cycle has not been rationally engineered so far, despite its major role in Corynebacterium glutamicum (6). From metabolic flux studies, however, it seems that the TCA cycle might offer a great potential for optimization (Fig. ​(Fig.1),1), which is also predicted from in silico pathway analysis (13, 22). Experimental evidence comes from studies with Brevibacterium flavum exhibiting increased lysine production due to an induced bottleneck toward the TCA cycle (21). In the present work, we performed TCA cycle engineering by downregulation of isocitrate dehydrogenase (ICD). ICD is the highest expressed TCA cycle enzyme in C. glutamicum (7). Downregulation was achieved by start codon exchange, controlling ICD expression on the level of translation.Open in a separate windowFIG. 1.Stoichiometric correlation of lysine yield (%), biomass yield (g/mol) and TCA cycle flux (%; entry flux through citrate synthase) determined by ¹³C metabolic flux analysis achieved by paraboloid fitting of the data set (parameters were determined with Y0 = 105.1, a = −1.27, b = 0.35, c = −9.35 × 10⁻³, d = −11.16 × 10⁻³). The data displayed represent values from 18 independent experiments with different C. glutamicum strains taken from previous studies (1-3, 11, 12, 15, 23).     

Contribution of the Periplasmic Chaperone Skp to Efficient Presentation of the Autotransporter IcsA on the Surface of Shigella flexneri

IcsA is an outer membrane protein in the autotransporter family that is required for Shigella flexneri pathogenesis. Following its secretion through the Sec translocon, IcsA is incorporated into the outer membrane in a process that depends on YaeT, a component of an outer membrane β-barrel insertion machinery. We investigated the role of the periplasmic chaperone Skp in IcsA maturation. Skp is required for the presentation of the mature amino terminus (alpha-domain) of IcsA on the bacterial surface and contributes to cell-to-cell spread of S. flexneri in cell culture. A mutation in skp does not prevent the insertion of the β-barrel into the outer membrane, suggesting that the primary role of Skp is the folding of the IcsA alpha-domain. In addition, the requirement for skp can be partially bypassed by disrupting icsP, an ortholog of Escherichia coli ompT, which encodes the protease that processes IcsA between the mature amino terminus and the β-barrel outer membrane anchor. These findings are consistent with a model in which Skp plays a critical role in the chaperoning of the alpha-domain of IcsA during transit through the periplasm.Type V secretion apparatuses (also called autotransporters) consist of an extensive class of large, outer membrane proteins of gram-negative bacteria, typically virulence factors, found in all subdivisions of proteobacteria (28). Although originally designated as “autotransporters” because they were thought to mediate their own insertion into and translocation across the outer membrane, more recent evidence suggests that autotransporter secretion and insertion requires the aid of accessory factors (21, 29). Secretion involves the insertion of the carboxy-terminal β-barrel domain into the outer membrane and translocation of the mature passenger (alpha) domain across the outer membrane (Fig. ​(Fig.1).1). Whether these two events occur sequentially or simultaneously is unclear. Analysis of crystal structures indicates that the carboxy-terminal end of the passenger domain is present within the central pore of the β-barrel (4, 27). Several studies provide evidence that at least some autotransporters are partially folded in the periplasm (7, 20), and one of these studies provides strong evidence that the passenger domain may be partially or fully incorporated into the β-barrel prior to incorporation of the mature protein into the outer membrane (20).Open in a separate windowFIG. 1.Schematic of the autotransporter IcsA. (A) Linear diagram showing the signal peptide (SP), alpha-domain (IcsA_53-757), and carboxy-terminal β-barrel domain. (B) IcsA in the outer membrane. The carboxy-terminal β-barrel is inserted into the outer membrane, and the mature amino-terminal alpha-domain is exposed on the bacterial surface. N′, mature amino terminus; C, carboxyl terminus; OM, outer membrane; arrow, proposed site of cleavage between residues 757 and 758 by IcsP.Shigella flexneri is a gram-negative human pathogen which, upon passage through the lower digestive tract, gains entry into colonic epithelial cells. Once S. flexneri is intracellular, it spreads to adjacent cells by secreting IcsA, a surface-associated autotransporter that is required for the polymerization of host cell actin on the bacterial surface. Actin polymerization occurs at a single pole of the bacterium and is required for infection of adjacent cells and disease pathogenesis (5, 24, 33).IcsA is encoded on a large virulence plasmid. The full-length protein is approximately 120 kDa and has three assigned functional and structural domains (25): an atypical Sec secretion signal (IcsA_1-52), the alpha-domain (IcsA_53-757), which is exposed on the bacterial surface and contains sequences that are required for actin polymerization, and the beta-domain (IcsA_758-1102), which forms a β-barrel structure in the outer membrane (Fig. ​(Fig.1A)1A) (21, 25). In vivo, a fraction of IcsA molecules are proteolytically processed at the junction between the alpha- and beta-domains by the protease IcsP (SopA), a protein which is also encoded on the virulence plasmid (14, 34). IcsA_53-757 is found in the supernatant of liquid cultures, while mature full-length IcsA (IcsA_53-1102), IcsA_758-1102 (14, 34), and some IcsA_53-757 (this work) remain cell associated. IcsA, like other autotransporters, is secreted at the bacterial pole (22), the site at which actin tail assembly occurs. As it is for other β-barrel-containing outer membrane proteins, insertion of IcsA and other autotransporters into the outer membrane requires the outer membrane insertase YaeT (BamA, Omp85) (21).Skp, DegP, and SurA are periplasmic chaperones that, like YaeT, appear to function in the targeting and/or insertion of outer membrane proteins (35). Evidence based on synthetic phenotypes suggests that during outer membrane protein insertion Skp and DegP act in one pathway and that SurA acts in a distinct but parallel pathway (35).We investigated the role of the periplasmic chaperone Skp in the folding and secretion of IcsA in S. flexneri. We found that in the absence of skp, IcsA is inefficiently presented on the surface of S. flexneri, leading to a cellular spread defect. Surprisingly, the protein was still efficiently cleaved by the outer membrane protease IcsP, as wild-type levels of IcsA_53-757 were detected in the culture supernatants. We found that introduction of the icsP mutation into the skp strain background led to an increase in the levels of full-length IcsA presented on the bacterial cell surface of the skp mutant, and we present models that could explain our results.     

Interactions of DNA with Biofilm-Derived Membrane Vesicles

The biofilm matrix contributes to the chemistry, structure, and function of biofilms. Biofilm-derived membrane vesicles (MVs) and DNA, both matrix components, demonstrated concentration-, pH-, and cation-dependent interactions. Furthermore, MV-DNA association influenced MV surface properties. This bears consequences for the reactivity and availability for interaction of matrix polymers and other constituents.The biofilm matrix contributes to the chemistry, structure, and function of biofilms and is crucial for the development of fundamental biofilm properties (46, 47). Early studies defined polysaccharides as the matrix component, but proteins, lipids, and nucleic acids are all now acknowledged as important contributors (7, 15). Indeed, DNA has emerged as a vital participant, fulfilling structural and functional roles (1, 5, 6, 19, 31, 34, 36, 41, 43, 44). The phosphodiester bond of DNA renders this polyanionic at a physiological pH, undoubtedly contributing to interactions with cations, humic substances, fine-dispersed minerals, and matrix entities (25, 41, 49).In addition to particulates such as flagella and pili, membrane vesicles (MVs) are also found within the matrices of gram-negative and mixed biofilms (3, 16, 40). MVs are multifunctional bilayered structures that bleb from the outer membranes of gram-negative bacteria (reviewed in references 4, 24, 27, 28, and 30) and are chemically heterogeneous, combining the known chemistries of the biofilm matrix. Examination of biofilm samples by transmission electron microscopy (TEM) has suggested that matrix material interacts with MVs (Fig. ​(Fig.1).1). Since MVs produced in planktonic culture have associated DNA (11, 12, 13, 20, 21, 30, 39, 48), could biofilm-derived MVs incorporate DNA (1, 39, 40, 44)?Open in a separate windowFIG. 1.Possible interactions between matrix polymers and particulate structures. Shown is an electron micrograph of a thin section through a P. aeruginosa PAO1 biofilm. During processing, some dehydration occurred, resulting in collapse of matrix material into fibrillate arrangements (black filled arrows). There is a suggestion of interactions occurring with particulate structures such as MVs (hollow white arrow) and flagella (filled white arrows) (identified by the appearance and cross-dimension of these highly ordered structures when viewed at high magnification), which was consistently observed with other embedded samples and also with whole-mount preparations of gently disrupted biofilms (data not shown). The scale bar represents 200 nm.     

A Putative ABC Transporter,HatABCDE, Is among Molecular Determinants of Pyomelanin Production in Pseudomonas aeruginosa

Pyomelanin overproduction is a common phenotype among Pseudomonas aeruginosa isolates recovered from cystic fibrosis and urinary tract infections. Its prevalence suggests that it contributes to the persistence of the producing microbial community, yet little is known about the mechanisms of its production. Using transposon mutagenesis, we identified factors that contribute to melanogenesis in a clinical isolate of P. aeruginosa. In addition to two enzymes already known to be involved in its biosynthesis (homogentisate dioxygenase and hydroxyphenylpyruvate dioxygenase), we identified 26 genes that encode regulatory, metabolic, transport, and hypothetical proteins that contribute to the production of homogentisic acid (HGA), the monomeric precursor of pyomelanin. One of these, PA14_57880, was independently identified four times and is predicted to encode the ATP-binding cassette of an ABC transporter homologous to proteins in Pseudomonas putida responsible for the extrusion of organic solvents from the cytosol. Quantification of HGA production by P. aeruginosa PA14 strains missing the predicted subcomponents of this transporter confirmed its role in HGA production: mutants unable to produce the ATP-binding cassette (PA14_57880) or the permease (PA14_57870) produced substantially less extracellular HGA after growth for 20 h than the parental strain. In these mutants, concurrent accumulation of intracellular HGA was observed. In addition, quantitative real-time PCR revealed that intracellular accumulation of HGA elicits upregulation of these transport genes. Based on their involvement in homogentisic acid transport, we rename the genes of this operon hatABCDE.Pseudomonas aeruginosa is a metabolically versatile, opportunistic pathogen that is a major cause of life-threatening infections in patients with burn wounds, compromised immunity, chronic obstructive pulmonary disease (COPD), and cystic fibrosis (CF) (23, 41). A major contributor to P. aeruginosa''s pathogenicity is its remarkable genomic plasticity, which often results is a wide range of phenotypic variation among isolates obtained from both acute and chronic infections. These phenotypes include small colony variant formation (24), alginate overproduction (36), hyperpigmentation (22), autoaggregation (13), and autolysis (64). Many of these phenotypes evolve as infections progress, and most have been ascribed to “loss-of-function” genome diversification that promotes long-term survival in the host environment (54). In this regard, recent studies have stimulated interest in another example of a loss-of-function phenotype, the mutation or deletion of hmgA, which encodes the homogentisate 1,2-dioxygenase enzyme. The absence of this protein leads to the accumulation and subsequent export of homogentisic acid (HGA), which ultimately aggregates into the pyomelanin polymer that manifests as a reddish brown pigmentation of P. aeruginosa colonies and their surrounding milieu (Fig. ​(Fig.1A)1A) (5, 49).Open in a separate windowFIG. 1.Pyomelanin production by the PA14 ΔhmgA and DKN343 strains. (A) Homogentisate pathway for the catabolism of chorismate and aromatic amino acids. Enzyme names are shown above the arrows for each step. A mutation or deletion of the hmgA gene (encoding homogentisate 1,2-dioxygenase) leads to the accumulation of pyomelanin. (B) Pyomelanin overproduction by the PA14 ΔhmgA mutant is abolished when complemented with an intact hmgA gene. Complementation of a melanogenic clinical P. aeruginosa isolate, DKN343, with hmgA results in no phenotypic change, indicating that other factors contribute to its pigmentation.Production of pyomelanin (and other forms of melanin) has been described to occur in a wide range of bacterial species, including Aeromonas (4), Azotobacter (51), Azospirillum (50), Bacillus (3), Legionella (8), Marinomonas (33), Micrococcus (40), Mycobacterium (45), Proteus (1), Rhizobium (12), Shewanella (61), Sinorhizobium (38), Streptomyces (67), and Vibrio (63) species. Notably, isolates of other bacterial species associated with chronic infections of the CF lung, Burkholderia cenocepacia and Stenotrophomonas maltophilia, can also be melanogenic (28, 58), suggesting a possible role for this pigment in the establishment and/or persistence of infection. Some genera produce melanin under normal conditions via polyphenol oxidases or laccases, while others synthesize the pigment only in response to specific environmental conditions (17, 35). Many species, however, including P. aeruginosa, overproduce pyomelanin as a result of a point mutation in hmgA or large chromosomal deletions of loci containing the homogentisate operon (2, 19). While these genetic variations have been frequently reported, there is little understanding of the competitive advantage, if any, that this pigment confers to the producing bacterium.Proposed roles for pyomelanin include the enhancement of bacterial surface attachment (20), extracellular electron transfer (61), iron reduction/acquisition (8), induction of virulence factor expression (63), heavy metal binding (21), and protection from environmental stress (11, 28, 32, 44, 53, 65). A protective role has also been proposed to occur in P. aeruginosa PA14, where pyomelanin was shown to contribute to the persistence of an overproducing strain in a chronic CF infection model in mice (49). However, given that melanogenic isolates have been recovered from laboratory-grown communities of P. aeruginosa PAO1 (5, 56), it is probable that pyomelanin plays other roles in addition to protection against host defense mechanisms.As a first step toward gaining a better understanding of pyomelanin function, we sought to identify the molecular determinants of its production in P. aeruginosa. By screening a library of pTnTet/minimariner transposon mutants of a pyomelanin-overproducing clinical isolate for alterations in pigmentation, we identified several genes whose products are involved in tyrosine catabolism, central metabolic pathways, nucleotide biosynthesis, regulation, and membrane transport, in addition to hypothetical proteins of unknown function. We chose to further characterize the gene identified most frequently in our screen, one annotated as encoding a putative ATP-binding cassette of an ABC-type transporter. Here, we demonstrate that this transporter is involved in HGA transport and the subsequent extracellular formation of pyomelanin.     

Bioproduction of p-Hydroxystyrene from Glucose by the Solvent-Tolerant Bacterium Pseudomonas putida S12 in a Two-Phase Water-Decanol Fermentation

Two solvent-tolerant Pseudomonas putida S12 strains, originally designed for phenol and p-coumarate production, were engineered for efficient production of p-hydroxystyrene from glucose. This was established by introduction of the genes pal and pdc encoding l-phenylalanine/l-tyrosine ammonia lyase and p-coumaric acid decarboxylase, respectively. These enzymes allow the conversion of the central metabolite l-tyrosine into p-hydroxystyrene, via p-coumarate. Degradation of the p-coumarate intermediate was prevented by inactivating the fcs gene encoding feruloyl-coenzyme A synthetase. The best-performing strain was selected and cultivated in the fed-batch mode, resulting in the formation of 4.5 mM p-hydroxystyrene at a yield of 6.7% (C-mol of p-hydroxystyrene per C-mol of glucose) and a maximum volumetric productivity of 0.4 mM h⁻¹. At this concentration, growth and production were completely halted due to the toxicity of p-hydroxystyrene. Product toxicity was overcome by the application of a second phase of 1-decanol to extract p-hydroxystyrene during fed-batch cultivation. This resulted in a twofold increase of the maximum volumetric productivity (0.75 mM h⁻¹) and a final total p-hydroxystyrene concentration of 21 mM, which is a fourfold improvement compared to the single-phase fed-batch cultivation. The final concentration of p-hydroxystyrene in the water phase was 1.2 mM, while a concentration of 147 mM (17.6 g liter⁻¹) was obtained in the 1-decanol phase. Thus, a P. putida S12 strain producing the low-value compound phenol was successfully altered for the production of the toxic value-added compound p-hydroxystyrene.The demand for so called “green” production of chemicals is rapidly increasing due to the declining availability of fossil fuels and the urgency to reduce CO₂ emissions (10, 30). However, this bioproduction may be hindered by the toxicity of the product of interest, such as substituted aromatics, to the production host (1, 2, 12, 29). One way to cope with this product toxicity is to deploy solvent-tolerant microorganisms as biocatalysts (5, 28). Of special interest among these solvent-tolerant hosts are Pseudomonas putida strains that have been engineered to produce a variety of compounds such as p-hydroxybenzoate (25, 33), p-coumarate (19), and (S)-styrene oxide (22). In our laboratory, we study and employ the solvent-tolerant P. putida S12. This strain is well suited for the production of substituted aromatic chemicals (18, 19, 33, 38) thanks to its extreme solvent tolerance (5, 35) and metabolic versatility toward aromatics (14, 16, 34).An example of an industrially relevant but extremely toxic aromatic is p-hydroxystyrene (4-vinyl phenol) (23). This compound is widely used as a monomer for the production of various polymers that are applied in resins, inks, elastomers, and coatings. Ben-Bassat et al. (2, 3, 23) reported p-hydroxystyrene production from glucose in Escherichia coli. In this strain, phenylalanine/tyrosine ammonia lyase (PAL/TAL; encoded by pal) from Rhodotorula glutinis and p-coumaric acid decarboxylase (PDC; encoded by pdc) from Lactobacillus plantarum were introduced for the conversion of l-tyrosine into p-hydroxystyrene via p-coumarate. The maximum concentration of p-hydroxystyrene was limited to 3.3 mM due to the toxicity of the product to the E. coli host (3, 23). To alleviate product toxicity, a two-phase fermentation with 2-undecanone as the extractant was performed. This approach resulted in a modest 14.2 mM p-hydroxystyrene in the organic phase and 0.5 mM p-hydroxystyrene in the water phase (2). Toxicity-related adverse effects on p-hydroxystyrene production may also be avoided by dividing the whole process into three stages: production of l-tyrosine from glucose by E. coli, conversion of l-tyrosine into p-coumarate by immobilized PAL-overexpressing E. coli cells, and chemical decarboxylation of p-coumarate into p-hydroxystyrene (29).In this report, we address and strongly enhance the bio-based production of p-hydroxystyrene from glucose by employing the solvent-tolerant P. putida S12 as a host. Previously, two strains, P. putida S12 C3 (19) and P. putida S12 TPL3 (38), have been constructed for the production of the l-tyrosine-derived aromatics p-coumarate and phenol, respectively. These strains were highly optimized for aromatics production, resulting in a heavily increased metabolic flux toward l-tyrosine. Therefore, they are suitable platform strains for the production of other l-tyrosine-derived aromatics (33). The bifunctional enzyme PAL/TAL (EC 4.3.1.25) from Rhodosporidium toruloides and the enzyme PDC (EC 4.1.1.-) from L. plantarum were introduced into these strains to allow the conversion of l-tyrosine into p-hydroxystyrene (Fig. ​(Fig.1).1). These minor modifications resulted in an efficient biocatalyst for the production of the value-added compound p-hydroxystyrene from glucose.Open in a separate windowFIG. 1.Schematic overview of the biochemical pathway for p-hydroxystyrene production. TAL, tyrosine ammonia lyase; FCS, feruloyl-coenzyme A synthetase. The cross indicates the disruption of fcs, disabling p-coumarate degradation.