The Aerobic and Anaerobic Respiratory Chain of <b><i>Escherichia coli</i></b> and <b><i>Salmonella enterica</i></b>: Enzymes and Energetics
GOTTFRIED UNDEN* AND PIA DÜNNWALD
[SECTION EDITOR: VALLEY STEWART]
Posted March 11, 2008
Institut für Mikrobiologie und Weinforschung, Johannes Gutenberg-Universität Mainz, 55099 Mainz, Germany
*Corresponding author. Mailing address: Institut für Mikrobiologie und Weinforschung, Johannes Gutenberg-Universität Mainz, Becherweg 15, 55099 Mainz, Germany. Phone: (49) 6131-3923550, Fax: (49) 6131-3922695, E-mail:
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Escherichia coli contains a versatile respiratory chain that oxidizes ten different electron donor substrates and transfers the electrons to terminal reductases or oxidases for the reduction of six different electron acceptors. Salmonella is able to use even two more electron acceptors. The variation is further increased by the presence of isoenzymes for some substrates. A large number of respiratory pathways can be established by combining the oxidation of different electron donors and acceptors. The respiratory dehydrogenases use quinones as the electron acceptors that are oxidized by the terminal reductase and oxidases. The enzymes vary largely with respect to their composition, architecture, membrane topology, and the mode of energy conservation. Most of the energy-conserving dehydrogenases (FdnGHI, HyaABC, HybCOAB, and others) and the terminal reductases (CydAB, NarGHI, and others) form a proton potential (▵p) by a redox-loop mechanism. Only two enzymes (NuoA-N and CyoABCD) couple the redox energy to proton translocation by proton pumping. A large number of dehydrogenases (e.g., Ndh, SdhABCD, and GlpD) and terminal reductases (e.g., FrdABCD and DmsABC) do not conserve the redox energy in a proton potential. For most of the respiratory enzymes the mechanism of proton potential generation is known from structural and biochemical studies, or it can be predicted from sequence information. The H+/2e− ratios of proton translocation for most respiratory chains are in the range from 2 to 6H+/e−. The energetics of the individual redox reactions and the respiratory chains is described and related to the H+/2e− ratios. In contrast to the knowledge on the function of enzymes and their role in proton potential formation, are the physiological aspects of respiration such as organization and coordination of the electron transport and the use of alternative respiratory enzymes not well characterized.
The chapter will present an overview on the respiratory enzymes and their role in respiration, both for well-characterized enzymes and for enzymes of which detailed knowledge is lacking. In addition, the bioenergetics of the alternative pathways and reactions will be presented. The overview continues and updates previous reviews on the respiratory chain of Escherichia coli (68, 216). Detailed presentation of selected respiratory enzymes will be given in chapters NADH as Donor, Respiration of Nitrate and Nitrite, Succinate as Donor; Fumarate as Acceptor, Oxygen as Acceptor, S- and N-Oxide Reductases, and Global Control of Respiratory Enzyme Synthesis. The respiratory chains of E. coli are composed each of two consecutive enzymes: the dehydrogenases functioning as quinone reductases and terminal reductases functioning as quinol oxidases. The dehydrogenases and terminal reductases are linked in this way by the quinones (Q), which can be ubiquinone (UQ), menaquinone (MK), or demethylmenaquinone (DMK). In this way branched respiratory chains are established where most of the dehydrogenases can interact with most of the terminal reductases. Three-membered, mitochondrial-type respiratory chains with quinol:cytochrome c reductase and cytochrome c reductase are not found in E. coli and other enteric bacteria. In E. coli no organization of respiratory enzymes in supercomplexes, as shown for mitochondria and some bacteria (184, 204), has been demonstrated so far and most of the enzymes might exist independently from each other.
In principle, most of the dehydrogenases are able to transfer electrons to each of the terminal reductases or oxidases, provided that the enzymes react with the same type of quinone. However, some of the dehydrogenases and terminal reductases are not expressed under the same conditions (see Global Control of Respiratory Enzyme Synthesis). Thus, many of the simple dehydrogenases, like NADH (Ndh), glycerol-3-P (GlpD), or D-lactate (DldD) dehydrogenases, are produced under aerobic conditions and, to a limited extent, in nitrate respiration, but they do not operate in anaerobic respiration with fumarate or dimethyl sulfoxide (DMSO). In nitrate respiration, the Ndh and GlpD enzymes therefore are used in parallel with the NuoA-N and GlpABC enzymes (22, 91, 194), but the "aerobic" enzymes Ndh and GlpD predominate (60). The activity of D-lactate dehydrogenase during nitrate respiration was also intermediate to the high and low activities of aerobic and fermentative growth, respectively (235).
The redox energy is conserved by redox-driven H+-pumping or redox half-loop enzymes. Enzymes using the Q-cycle for driving proton potential formation are not found In E. coli. This type of redox-driven proton translocation is verified in quinol-cytochrome c oxidoreductase (cytochrome bc1 complex) from mitochondria and bacteria like Paracoccus are not found in E. coli (25, 90). Redox-driven H+-pumping has been shown for NADH dehydrogenase I (NuoA-N) and cytochrome oxidase bo3 (CyoABCD). The redox-loop mechanism is of particular significance for energy conservation in E. coli and related bacteria. This mechanism was suggested originally by P. Mitchell as the main principle for driving proton potential formation by respiration (155). In the redox loop two electrons are transferred from an electron donor at the positive side of the membrane to the negative side where a quinone is reduced. The reduction of the quinone consumes two protons at the same time. For the second half-loop, the quinol crosses the membrane to the positive side where it is oxidized and releases protons. In this way, the complete redox loop generates a proton potential by the transfer of electrons across the membrane and uptake and release of protons on opposite sites of the membrane. The redox loop is catalyzed by two enzymes that catalyze a half-loop each. This type of energy conservation is found in (anaerobic) formate dehydrogenase FdnGHI, nitrate reductase NarGHI, and other enzymes (see Fig. 8). A considerable number of enzymes, such as succinate dehydrogenase SdhABCD, fumarate reductase FrdABCD, and the one-subunit peripheral membrane enzymes like NADH dehydrogenase II (Ndh) or D-lactate dehydrogenase (DldD), release and consume H+ on the same side of the membrane (scalar H+) and do not contribute directly to the generation of proton potential (or ▵p). Their significance lies in the supply or consumption of electrons to enable the function of the second respiratory enzyme of the electron transfer chain which is then a coupling enzyme that conserves the free energy of the redox reaction in a proton potential, either by pumping H+ across the membrane, or by the use of a redox half-loop mechanism.
In E. coli 15 different primary dehydrogenases have been described biochemically and genetically (Fig. 1A to D and Fig. 1E). In addition, some dehydrogenases of biosynthetic and other pathways link anabolic redox reactions to the quinones and constitute "anabolic respiratory chains" for disposal of electrons (Fig. 4).
Fig. 1A to D and Fig. 1E group the respiratory dehydrogenases according to structural, functional, and topological aspects. Principally the following groups can be identified: redox-driven H+ pumps that translocate H+ as a consequence of the redox reaction (Fig. 1A); composite enzymes with a redox half-loop, transferring electrons across the membrane due to the transmembrane topology of the active sites for their substrate and the quinone, and generating a proton potential by the topology of the active sites for their substrate and for the quinone (Fig. 1B); composite enzymes without a redox half-loop with both active sites on the same side of the membrane (Fig. 1C); small enzymes consisting of one subunit that carry the active sites for both substrates on the same side of the membrane (Fig. 1D); and GlpABC, which is a composite enzyme differing from the composite enzymes of types B and C (Fig. 1E). Enzymes of types C and D do not form a H+ potential since neither electrons nor protons pass the membrane during the reaction. Dehydrogenases of the latter types conserve redox energy only in combination with subsequent energy-conserving steps.
NADH dehydrogenase I (NADH: quinone reductase, or NuoA-N) (Fig. 1A) appears to be the only H+-pumping dehydrogenase of the respiratory chain of E. coli (for a detailed review on NADH dehydrogenase I, see chapter NADH as Donor and reference 61). NADH dehydrogenase I (NADH + H+ + Q + 4H+in → NAD+ + QH2 + 4H+ex) utilizes NADH and deamino-NADH (dNADH) as a substrate, which allows differentiation from NADH dehydrogenase II reacting only with NADH. The enzyme consists of 13 subunits and is homologous to the H+-pumping NADH dehydrogenase I of mitochondria and other respiratory bacteria (61, 79, 135, 226, 231). The enzyme contains all prosthetic groups and subunits of the mitochondrial enzyme that have a function in catalysis and electron transfer. Accessory subunits for structural and regulatory functions of the mitochondrial enzymes are missing, and the enzyme is regarded as a minimal form of a H+-pumping NADH-dehydrogenase I.
NADH dehydrogenase I functions in aerobic and anaerobic respiration (23). In aerobic growth the noncoupling NADH dehydrogenase II represents the major enzyme, whereas in fumarate or DMSO respiration NADH dehydrogenase I is essential (28, 29, 210). The enzyme contains a large peripheral arm in the cytosol comprising six subunits. This subcomplex carries the site for NADH oxidation (with bound flavin mononucleotide [FMN]) and forms ligands for nine FeS clusters of the [2Fe-2S] and [4Fe-4S] types that transfer the electrons from the NADH site to the membrane arm. The structure of the peripheral complex of NADH dehydrogenase I from Thermus thermophilus has been solved by X-ray crystallography (183). The FMN and seven of the FeS clusters are linearly arranged with edge-to-edge distances of ≤14 Å in accordance with a linear electron transfer across the enzyme. The membrane integral part of the enzyme consists of seven hydrophobic subunits. It contains the site for quinone reduction and is supposed to translocate the protons in an unknown mode (61, 88, 115, 134). NADH dehydrogenase I appears to be a redox-driven H+ pump (202). The enzyme is also able to drive Na+ translocation (198, 199), which could depend on the function of a secondary Na+/H+ antiport activity. The enzyme represents one of the most important primary dehydrogenases in the respiratory chain of E. coli and is described in detail in chapter NADH as Donor.
Formate dehydrogenase FdnGHI, and most likely also formate dehydrogenase FdoGHI and hydrogenases HyaABC and HybCOAB, conserve the free energy of the redox reactions by a redox-loop mechanism (Fig. 1B).
E. coli contains three formate dehydrogenases. Two of the formate dehydrogenases (FdnGHI and FdoGHI) are membrane integral and linked to respiration. The third enzyme (Fdh-H) is part of the formate-hydrogen-lyase complex (FHL) and of a similar complex (FHL-2) encoded by the hyf operon with a supposedly similar function. Formate dehydrogenase FdnGHI is active under anaerobic conditions and functions mainly in nitrate or nitrite respiration (13, 47, 59, 137). Formate is provided either from the medium or as a product of mixed (or formic) acid fermentation. The enzyme shows a "mushroom"-like overall structure (109, 110) that is characteristic for composed respiratory enzymes that react with a hydrophilic and a hydrophobic substrate (Fig. 1B and C). The structure consists of a stalk formed by the membrane anchor (FdnI) and of a head composed of the hydrophilic catalytic (FdnG) and the electron transfer (FdnH) subunits (Fig. 1B). FdnH contains, in addition, a hydrophobic membrane-embedded α-helix and with the four transmembrane helices of FdnI forms the membrane anchor of five α-helices. The formate site of Fdn is in the periplasm. Quinones serve as the electron acceptors of the reaction (HCO2− + H2O + Q + 2H+in → HCO3− + QH2 + 2H+ex). The reaction includes transfer of two electrons from the formate to the quinone site, and the consumption and release of 2H+ at the cytoplasmic and periplasmic sites, respectively. Formate oxidation is catalyzed at a molybdopterin-guanine dinucleotide cofactor (MGD) in FdnG. The site for quinone reduction is on subunit FdnI in the membrane close to the cytoplasmic side. The electron transfer from the MGD to FdnI is mediated by five [4Fe-4S] clusters that are located in FdnG and FdnH. The electrons are transferred to the periplasmic-oriented heme b group (heme bP) in FdnI close to the periplasmic side of the membrane, and then to the second heme bC (see Fig. 8). Heme bC is close to the cytoplasmic side of the membrane and represents the site for quinone reduction. The edge-to-edge distances of the redox cofactors are a maximum of 10.7 Å, which allows rapid electron transfer. Only the role of the first FeS cluster in electron transfer is not clear. The location and topology of the enzyme and of the active sites for formate and Q are well characterized (110), proving the function of the enzyme in a redox loop and in H+ translocation (108).
Formate dehydrogenase FdoGHI also catalyzes formate:Q reduction (1, 167). The enzyme is very similar in sequence and composition to FdnGHI. FdoGHI shows low enzymatic activity and is produced under aerobic conditions. The enzyme appears to be part of an aerobic formate–nitrate respiratory chain. Very little is known about the enzyme and aerobic formate oxidation by respiration. Subunit composition and prosthetic groups appear to be very similar to FdnGHI, suggesting a similar overall structure and function. The topology of Fdo is not known. It has been suggested that is the inverse to FdnGHI (12) with FdoG, FdoH, and the formate site in the cytoplasm. The topology derived from the studies might be preliminary and complicated by the involvement of the Tat protein transport pathway (197). Due to the complications it is assumed in Fig. 1A to D and Fig. 1E (without experimental evidence) that FdoGHI has the same topology as FdnGHI for reasons of similarity. FdoI contains four conserved His residues, suggesting binding of two heme b residues similar to FdnI, and catalysis of transmembrane electron transfer.
E. coli contains four different hydrogenases chapter (Anaerobic Formate and Hydrogen Metabolism). Two of the hydrogenases (Hyd-1 and Hyd-2) are (respiratory) uptake hydrogenases, whereas the other two (Hyd-3 and Hyd-4) are part of formate-hydrogen-lyase complexes FHL and FHL-2. The hydrogenases are [NiFe]-hydrogenases with a Ni-Fe cofactor at the active site. The uptake hydrogenases HyaABC (Hyd-1) and HybCOAB (Hyd-2) link oxidation of H2 to the reduction of quinones and the respiratory chain chapter (Anaerobic Formate and Hydrogen Metabolism). Both hydrogenases have periplasmic orientation. The enzymes have been used, together with the hydrogenase 3 of formate hydrogen-lyase FHL, to study molecular biology of hydrogenase and NiFeS-cofactor maturation (for reviews, see reference 21). Structural and functional aspects of respiratory dehydrogenases have been studied in more detail in Wolinella succinogenes, which contains an energy-conserving hydrogenase HydBACWs homologous to HyaABC of E. coli (52, 74, 76, 77, 215). We will refer to this enzyme for structural and functional aspects.
Hydrogenase 1 (HyaABC) is produced under anaerobic conditions and supposed to function in H2 reoxidation, which has been produced from formate by formate hydrogen lyase (FHL) (151, 152, 172, 179, 181, 182). The enzyme is similar in sequence and composition to hydrogenase HydBACWs, which has periplasmic orientation and functions as a hydrogen-Q reductase (52, 75). The membrane anchor HydCWs (homologous to HyaC of E. coli) is a hydrophobic protein with four predicted TM helices (74, 76, 77). HydCWs binds two heme b groups in transmembrane arrangement, one close to the periplasmic and the other close to the cytoplasmic side. The four His ligands of the heme b groups are found on three of the TM helices (76). The active site for MK reduction is close to the periplasmic heme b (74). HydBWs, the NiFeS-containing catalytic (large) subunit, together with HydAWs, forms the hydrophilic head of the enzyme. HydAWs carries FeS clusters for electron transfer from the catalytic subunit to HydCWs. HydAWs contains (like FdnH) a hydrophobic α-helix that anchors the subunit together with HydC directly in the membrane, resulting in a five-α-helical membrane anchor for the HydBAC trimer (74, 77). HydBACWs has all the structural und functional prerequisites for a redox-loop enzyme. In H2-fumarate respiration of W. succinogenes the enzyme is the only site for generation of the proton potential (126), and it can be assumed that the redox loop supplies the mechanism for generation of a proton potential. HyaABCEc, which is similar to HydBACWs in sequence and function, presumably shows the same architecture and applies the same mechanism for generation of a proton potential.
Hydrogenase 2 (HybCOAB or Hyd-2) of E. coli has been studied by molecular biological and genetic methods (9, 53, 151, 178). It is suggested that hydrogenase 2 is part of the electron transport from H2 to fumarate. The reaction allows growth by fumarate respiration (16). In H2-fumarate respiration of E. coli hydrogenase HybCOAB is the only energy-conserving enzyme and has to conserve redox energy in a proton potential. HybCO represents a soluble fragment of the enzyme and oxidizes H2 with benzyl viologen as the acceptor (9, 53). HybC is the large catalytic subunit with a [NiFe] binding site. The electron transfer subunit HybO contains three Cys clusters for the binding of FeS centers. HybA is a soluble protein of unknown function. It is predicted to contain two [4Fe-4S] clusters and might function in electron transfer (178). HybB represents the membrane anchor of the protein. HybB is significantly longer than HyaC and has about twice the number of transmembrane helices of HyaBC (10 to 11). The transmembrane α-helices carry four conserved His residues that presumably serve as the ligands for two heme b molecules, similar to HydCWs and HyaCEc (77, 151). The electron acceptor of Hyd-2 has not been identified experimentally (133), but MK has been supposed as the acceptor since the MK-dependent fumarate reductase is the final acceptor. Since H2-fumarate respiration supports growth of E. coli (16), Hyd-2 should be capable of energy conservation, presumably by a redox-loop mechanism similar to HyaABC and HydBACWs.
The only dehydrogenase of this type in E. coli is succinate dehydrogenase linking succinate oxidation to quinone reduction (succinate + Q → fumarate + QH2) (Fig. 1C). The structure of the enzyme (SdhABCD) has been solved by X-ray crystallography (236). For a detailed description and references, see chapter Succinate as Donor; Fumarate as Acceptor and references 32, 130, and 209. SdhC and SdhD provide the membrane anchor with six transmembrane helices. SdhA and SdhB are fixed to the membrane by protein contacts to SdhCD. The site for succinate oxidation is on SdhA with a covalently bound FAD. The electrons are transferred to the membrane anchor by three FeS clusters in SdhB ([2Fe2S] → [4Fe4S] → [3Fe4S]). SdhCD binds a heme b group by two His ligands at a site close to the [3Fe-4S] cluster of SdhB. The His ligands are supplied from either subunit. The active site for Q reduction is close to the heme b and formed from SdhCD and SdhB.
By this arrangement, H+ release from succinate and H+ uptake during Q reduction occur at the same side of the membrane, resulting in no charge or H+ translocation by the overall reaction. The reaction is not electrogenic and does not contribute to proton potential formation. Succinate dehydrogenase from bacteria containing only MK like Bacillus and Desulfovibrio (185, 237) carry a second heme b close to the opposite (outer) side of the membrane, which is the site for MK reduction (81). Consequently, protons are consumed at the outer side of the membrane. The (endergonic) reduction of MK by this type of succinate dehydrogenase is electrogenic and represents a reverse redox loop that consumes a membrane potential to drive the redox reaction (185, 187, 237). The architecture of E. coli SdhCDAB is similar to redox-loop enzymes, but lacks transmembrane electron transfer, suggesting that the enzyme is derived from a redox-loop enzyme, or is an evolutionary precursor of this type of enzyme.
Dehydrogenases consisting of one subunit represent the type of dehydrogenase occurring most frequently in the respiratory chain of E. coli (Fig. 1D). The enzymes are tightly membrane bound or only membrane associated (Fig. 2). The sites for the reaction of the electron donor substrate and of the quinones are close to each other. Dehydrogenases of this type are generally devoid of redox groups for the transfer of electrons from the substrate to the quinol site. Protons are released and consumed at the same side of the membrane (scalar protons), and none of the enzymes generates a proton potential directly. The dehydrogenases of this type mostly contain flavin adenine dinucleotide (FAD), FMN, or pyrroloquinoline quinone (PQQ) as a redox cofactor at the substrate site that is reduced by the primary donor and reoxidized by the quinones. The FAD and PQQ cofactors therefore are close to the membrane side to enable reaction with the quinones.
Membrane-bound enzymes.
Glucose dehydrogenase (Gcd) (glucose H2O + Q → gluconate + QH2) (Fig. 1D) is produced under aerobic conditions and provides an alternative mode to (normal) glucose oxidation by glycolysis (4, 89, 232, 234). The physiological role of the enzyme is not clear. Gcd is permanently bound to the membrane and has a periplasmic orientation, which is unique among the one-subunit dehydrogenases of E. coli. The enzyme contains a PQQ prosthetic group at the glucose site (45). PQQ cannot be produced by E. coli and has to be supplied by the medium to form the functional enzyme (219).
The protein has two distinct domains (Fig. 2): an N-terminal hydrophobic domain (amino acids [aa] 1 to 150) and the C-terminal domain (aa 151 to 796) with the conserved PQQ binding site (45, 232, 233). Membrane binding and insertion occurs by five transmembrane α-helices of the N-terminal domain. The C-terminal domain with the site for glucose oxidation and PQQ binding extends into the periplasm. The site for Q reduction appears to be located in the C-terminal domain as well, possibly at the membrane/protein interface close to the PQQ site (57), but bioinformatic studies indicate also that the Q site resides in the N-terminal anchor domain (62).
Membrane-associated (or peripheral) enzymes.
Most of the simple dehydrogenases are membrane associated (or peripheral) and do not contain transmembrane α-helices (Fig. 2). The proteins bind to the membrane by hydrophobic and/or electrostatic interaction. Proteins bound by electrostatic forces display an electropositive surface and interact mainly on the membrane surface with the electronegative phospholipid head groups. The hydrophobic interactions can be via apolar α-helices or loops on the protein surface that penetrate into a single leaflet of the membrane.
E. coli is able to use D-lactate as a carbon and energy source for aerobic growth. D-Lactate is oxidized by D-lactate dehydrogenase (Dld, dld gene) (30, 113, 143, 176, 207). D-Lactate dehydrogenase is a peripheral membrane protein with cytoplasmic orientation (Fig. 1D and Fig. 2). The enzyme has no particular polar segments and can be washed from the membrane with guanidine hydrochloride, but detergent is required for purification. The crystal structure of the enzyme has been solved (54). It consists of three domains: a FAD binding (catalytic) domain, a cap domain, and the membrane-association domain. The noncovalently bound FAD is at the catalytic site for D-lactate oxidation, which is covered on the membrane distal side by the cap domain. The membrane-associated part of the protein is rich in basic (Lys, Arg) amino acid residues, which are supposed to bind by electrostatic interactions to the negatively charged phospholipid head groups on the membrane surface. In addition, a region of the protein rich in hydrophobic (Tyr, Phe) residues was suggested to interact with the upper leaflet of the lipid layer (206). In this way, the enzyme is attached to the surface of the membrane without deeply penetrating the membrane.
The enzyme is supposed to donate electrons directly from the FADH2 to the membrane integral quinones. The reactive head groups of the quinones are located close to the membrane surface or are accessible from the peripheral part of the membrane (54). The site of Q binding and interaction with Dld has not been identified, but its location is expected in the membrane attachment site that allows close accession of the quinones to the FAD. According to this assumption all parts of the reaction occur close to the cytoplasmic surface of the membrane.
Pyruvate:Q reductase (PoxB, poxB gene) (pyruvate H2O + Q → acetate + CO2 + QH2) catalyzes the oxidative decarboxylation of pyruvate, using Q as the electron acceptor (36, 67). The enzyme (sometimes termed pyruvate oxidase) therefore functions as a pyruvate:Q reductase (or dehydrogenase). Decarboxylation requires thiamine diphosphate (Mg2+ TPP) as a cofactor. The enzyme is a homotetramer of the 62-kDa monomer (72, 223). Each of the monomers contains a tightly (noncovalently) bound FAD. Similar to other peripheral membrane enzymes, PoxB is activated by detergent and/or phospholipid, which are suggested to bind to the C-terminal part of the protein (34, 35). The C-terminal part represents the supposed membrane binding site. In bacteria the enzyme is inactive and located in the cytosol at low pyruvate concentrations (37, 67). High pyruvate concentrations trigger a conformational change that exposes the C-terminal lipid binding domain (31, 123). The protein then attaches to or inserts into the membrane, which increases the activity of the enzyme by releasing an inhibitory effect of the C-terminal part. In this way the FADH2 of the enzyme gains access to the membrane and the quinones that accept the electrons.
Enzymes with an unknown form of membrane association.
NADH dehydrogenase II (Ndh, ndh gene) represents an alternative NADH:quinone reductase encoded by the ndh gene (104, 105, 144). The enzyme (45 kDa) is of much simpler composition than the NuoA-N isoenzyme. Ndh contains noncovalently bound FAD. The enzyme is produced under aerobic conditions and apparently not used in anaerobic respiration (196, 210). In aerobic growth the enzyme is used in parallel with NuoA-N. Ndh binds in an unknown way to the membrane, presumably by a predictable hydrophobic transmembrane α-helix (19). The enzyme donates the electrons from NADH via FAD to the respiratory quinones, and for isolated Ndh high enzyme activities are found only when hydrophobic quinones are used, suggesting that the active site for the quinone is accessible from the membrane (19).
Mqo (originally termed malate oxidase) is a peripheral malate:Q reductase (malate + Q → oxaloacetate + QH2), which oxidizes endogenously produced malate (from citric acid cycle) or malate taken up from the external medium (157, 218). Mqo and the (classical) soluble NADH-dependent malate dehydrogenase (MDH) operate at the same time in E. coli. MDH appears to be the major enzyme under most conditions, and Mqo was synthesized only during exponential growth. Deletion of mqo has, in contrast to the deletion of the mdh gene, no clear effect on growth and metabolism under most conditions. The physiological role of the Mqo enzyme is not clear, therefore. In Corynebacterium glutamicum, however, the enzyme is the principal enzyme for malate oxidation (156). Because of the differences in the redox potential of the acceptors (−320 mV for NAD+, and +113 or −74 mV for Q or MK), it was suggested that use of Mqo and of the quinones might be favorable for malate oxidation, whereas MDH and NADH should be used preferentially for oxaloacetate reduction. This suggestion could not be verified for E. coli.
Membrane binding of Mqo and reaction with the quinones has not been studied in detail. The enzyme contains no predicted hydrophobic transmembrane α-helix. The enzyme can be detached from the membrane by sonication or membrane disintegration, and the activity is stimulated by detergents and phospholipids.
L
-Lactate dehydrogenase (LldD, lldD or lctD gene) is an inducible L-lactate specific dehydrogenase of 42.7 kDa that is used mainly in aerobic metabolism (51, 64, 96). The enzyme is membrane bound and donates the electrons of L-lactate to the quinones (L-lactate + Q → pyruvate + QH2). The enzyme contains FMN as a prosthetic group. It requires detergent for solubilization and oligomerizes in the absence of detergent (119). LldD contains no predicted transmembrane helices. Details on the biochemistry and function of the enzyme are not known.
Glycerol-3-P dehydrogenase (GlpD, or glp gene) is used for oxidation of glycerol-3-P in aerobically growing E. coli. GlpD is a homodimeric flavoprotein (GlpD2) and lacks a predictable hydrophobic transmembrane α-helix (7, 189). The enzyme is fully active only when membrane inserted and it donates electrons to quinones (glycerol-3-P + Q → dihydroxy-acetone-P + QH2). It is suggested that the enzyme is inserted by an amphipathic helix in the hydrophobic core region of the membrane (222).
D-Amino acid dehydrogenase (DadA) oxidatively deaminates D-amino acids to the corresponding oxoacid (103, 140). DadA is a membrane-associated flavoprotein and couples the reaction to the reduction of quinone (163). D-Ala is used with the highest preference (D-Ala + H2O + Q → pyruvate + NH3 + QH2). The protein was solubilized from membranes and purified by using detergent (107, 163). The enzyme preparation contains two subunits (55 kDa and 45 kDa), but only the gene for the smaller subunit was identified (dadA gene) (103, 140). Details on membrane binding and on the quinone site are not known.
GlpABC (glpABC genes) is the anaerobically expressed glycerol-3-P dehydrogenase that is used in anaerobic respiration (42, 97, 120, 128, 220). Operon and enzyme are named alternatively glpACB and GlpACB (97, 120, 220), but here, the terminology according to glpABC and GlpABC is used. The enzyme (Fig. 1E) is suggested to couple the oxidation of glycerol-3-P to fumarate, but also to nitrate respiration. The enzyme is very labile and has not been studied biochemically. The putative catalytic dimer GlpAB is composed of the hydrophilic proteins GlpA and GlpB of 62 kDa and 43 kDa (188). The sequence suggests that GlpA and GlpB bind FAD and FMN, respectively, as cofactors. GlpC is associated with the membrane and presumably functions as the membrane anchor for GlpAB (220). The sequence predicts for GlpC amphipathic helices, which are supposed to anchor the protein in the membrane (42) (Fig. 1E). Details on the membrane insertion of GlpC, and whether the protein is a transmembrane, are not known. GlpC contains two cysteine clusters typical for FeS-binding proteins, and EPR spectra indicate the presence of FeS clusters. By the lack of transmembrane helices and the presence of FeS clusters in the membrane anchor, the protein differs from other respiratory enzymes of E. coli. The coupling of GlpABC to the menaquinone-dependent fumarate reductase FrdABCD suggests that menaquinone is the direct electron acceptor of glycerol-3-P oxidation. It is also not clear whether the enzyme couples the reaction to the formation of a proton potential. The low redox potential difference for electron transfer from glycerol-3-P to menaquinone (▵E0' = 116 mV) argues against formation of a proton potential by the enzyme (see Table 1). GlpABC differs, therefore, by the type of membrane anchor and the redox group in the membrane anchor from other membrane-bound primary dehydrogenases of the respiratory chain.
Table 1Energetic parameters for the respiratory dehydrogenases of E. coli and S. enterica serovar Typhimurium |
E. coli contains membrane-bound enzyme complexes that are composed of two redox enzymes. Formate hydrogen-lyase (FHL) consists of a formate dehydrogenase (FDH-H) and a hydrogenase (Hyc enzyme) (Fig. 3; for review, see chapter Anaerobic Formate and Hydrogen Metabolism and references 136 and 180). Hydrogenase (HycBFEG) and formate dehydrogenase (FDH-H) are loosely bound to the HycCD proteins, which are membrane integrated and comprise 20 to 22 transmembrane helices. The Hyc and FDH-H proteins form a labile membrane-associated complex that transfers electrons from formate to protons, resulting in H2 formation. Subunits HycBFG contain clusters of Cys residues typical for binding of FeS clusters and are supposed to transfer electrons from the catalytic formate dehydrogenase subunit FDH-H to the catalytic hydrogenase subunit HydE. The reaction is reversible (HCOO− + H+ ↔ CO2 + H2), and the free-energy ▵G0' of the reaction is close to 0 kJ/mol formate under standard conditions. At high formate concentrations or acidic pH the conversion of formate to CO2 + H2 is energetically more favorable (−5.7 kJ/mol formate or pH unit) and becomes exergonic (up to −20 kJ/mol). Subunits of HycCDEFG share sequence similarity to subunits of NADH dehydrogenase I (Nuo). It has been suggested that formate hydrogen-lyase drives proton translocation, but there is no experimental proof for this.
A gene cluster (hyf genes) homologous to the genes of the FHL complex has been identified (6). The hyf genes code for a putative complex HyfABCDFGHI, which is homologous to the HycBCDEFG proteins of FHL. The HyfB, D, and F proteins correspond to HycC, which results in composition of the (putative) Hyf complex from eight Hyf proteins compared with the six Hyc subunits in FHL. The hyf cluster encodes, in addition, a membrane protein (HyfE) of unknown function that has no counterpart in FHL. It was suggested that the Hyf complex forms a proton-translocating formate-hydrogenlyase FHL-2 together with FDH-H (6), but the energetic limitations for proton translocation are the same as mentioned previously for FHL. The expression of the hyf operon is very weak, and the conditions for the synthesis of FHL-2 and its physiological function are not known (190, 194).
The membrane-bound RsxACDGE complex of E. coli functions together with the RscE gene product as a reducing system for the [2Fe-2S] cluster of the superoxide sensor SoxR. SoxR is converted by superoxide and NO to the oxidized state which then activates the expression of the target gene soxS (203). For reversion to the inactive state the FeS cluster of SoxR has to be reduced by the RsxACDGE complex by using NADPH as the electron donor (NADPH-dependent SoxR reductase) (122, 124). RsxACDGE is highly homologous to the RnfABCDGE complex of Rhodobacter capsulatus, which is thought to transfer electrons from NADPH to nitrogenase (111, 127, 186). For this reason the Rsx complex of E. coli was renamed in databases as RnfABCDGE. In Clostridium tetanomorphum a homologous enzyme system is found that functions as a membrane-bound NADH-ferredoxin-oxidoreductase, which is part of the glutamate fermentation pathway (22). The NADH-Fd-oxidoreductase is supposed to couple the exergonic reduction of NAD by ferredoxin to Na+ translocation across the membrane.
The redox potential of SoxR has been estimated to be −285 mV (50), suggesting rapid reduction by NAD(P)H, which has an estimated redox potential of −340 mV under cellular conditions (124). The RsxACDGE complex is predicted to be membrane bound by the membrane integral proteins RsxADE. In addition, RsxB and RsxG are supposed to be membrane associated in the complex by single transmembrane loops. The significance of membrane binding for function of the Rsx complex is not known.
Oxidation reactions of biosynthetic pathways are normally catalyzed by NAD(P)-dependent soluble dehydrogenases or oxidases. Some Q-linked dehydrogenases of biosynthetic reactions, however, link the reactions to the respiratory chain (Fig. 4).
Dihydroorotate dehydrogenase (PyrD) (also termed dihydroorotate oxidase) catalyzes the oxidation of dihydroorotate to orotate in the biosynthesis of pyrimidine ribonucleotides (5, 116). PyrD works under aerobic and anaerobic conditions and transfers the electrons to ubiquinone or menaquinone (dihydroorotate + Q → orotate + QH2). The final acceptors are oxygen, fumarate, or nitrate, and presumably other quinol-oxidizing terminal reductases. PyrD (37 kDa) is a homodimer (PyrD2) and contains FMN as the cofactor (117, 132). PyrD is a class 2 dihydroorotate dehydrogenase that is membrane associated and uses quinones as the acceptors (116). Class 2 dihydroorotate dehydrogenases are found in gram-negative bacteria like E. coli and in the mitochondria of most eucaryotes. The N terminus of the enzyme is arranged in three hydrophobic α-helices (αA, αB, and αC), which are supposed to attach the protein to the membrane (160). The helices contain also the active site for the quinone. The catalytic FMN is located close to the dihydroorotate and the quinone sites. It is accessible for dihydroorotate from the cytoplasmic, and for the quinones from the membrane space. During catalysis, FMN is first reduced by the dihydroorotate, and the FMNH2 then transfers the electrons to the nearby quinone.
In E. coli periplasmic thiol-disulfide oxidoreductases (Dsb enzymes) facilitate disulfide bond formation of periplasmic proteins for proper folding and stability. The periplasmic Dsb proteins, DsbA and DsbB, function as protein disulfide reductases, catalyzing protein thiol oxidation by the membranous quinones. DbsA directly oxidizes Cys thiols of secreted proteins and introduces disulfide bonds. DbsB reoxidizes DbsA by using quinones as the acceptors (2 Protein-SH + Q → protein-disulfide + QH2) (8). DsbB is membrane integral with four TM helices and forms a complex with the peripheral protein DsbA (93). DsbA reduces two Cys residues (Cys41 and Cys44) of DsbB, which are then oxidized by the quinone, resulting in a Cys41-Cys44 disulfide. Cys41 and Cys44 are close to transmembrane helix 2 of DsbB. The residues are accessible from the membranous quinone and from Cys130 of DsbB which transfers the electrons from DsbA according to a present model (93).
The sensor kinase ArcB of the two-component sensor system ArcBA responds to the presence of O2 via the redox state of the quinones. ArcB is anchored by two transmembrane helices in the membrane (142). The cytoplasmic part of the ArcB protein contains two Cys residues (Cys180 and Cys241) close to the cytoplasmic side of the membrane. In the presence of O2 the residues are oxidized to intermolecular disulfides (Cys180)2 and (Cys241)2 causing inactivation of the kinase activity. The oxidation of the Cys-thiols is effected by the respiratory quinones in a reaction similar to that of DsbB. ArcB is supposed to become oxidized specifically only by UQ, whereas DsbB and PyrD use UQ and MK as the electron acceptor.
Protoporphyrinogen "oxidase" (hemG product) catalyzes the oxidation of protoporphyrinogen IX to protoporphyrin IX in porphyrin or heme biosynthesis (101, 159). The enzyme is presumably a Q-dependent dehydrogenase that transfers the electrons to the respiratory quinones (protoporphyrinogen IX + 3 Q → protoporphyrin IX + 3 QH2) (46). The reaction takes place in aerobically and anaerobically growing bacteria. Details of enzyme function and on its interaction with the membrane and the quinones are not known.
E. coli and Salmonella contain at least 14 different terminal membrane-bound reductases (Fig. 5A to C, Fig. 5D, and Fig. 6) for the reduction of altogether eight different electron acceptors (O2, nitrate, nitrite, fumarate, trimethylamine N-oxide [TMAO], DMSO, tetrathionate, and thiosulfate). Terminal reductases consisting of only one subunit are missing. For O2 two, and for nitrate, three different terminal reductases are available. The expression of the structural genes is subject to transcriptional regulation by O2 and other electron acceptors (see Global Control of Respiratory Enzyme Synthesis ). All of the enzymes use reduced quinones as the electron donors. Some of the enzymes function as H+ pumps or as redox half-loops, whereas others constitute electron sinks without generating a proton potential directly. For quinol oxidase CyoABCD, nitrate reductase NarGHI, and fumarate reductase FrdABCD, detailed structural and biochemical information is available that allows understanding of the enzymes and their reactions and of the H+/e− ratios at the molecular level. For other enzymes many functional aspects can be derived from the function of analogous enzymes.
Quinol oxidase CyoABCD (or quionol oxidase bo3) is a member of the heme Cu oxidase family that couples the redox reaction to H+ pumping (2H+/2e−) (228). In addition 2H+/2e− are "translocated" by the release of 2H+ during quinol oxidation at a side close to the periplasm, and the consumption of 2H+, resulting in an overall ratio of 4H+/2 e− (Fig. 5A to C and Fig. 5D). CyoABCD is produced under aerobic conditions, whereas the alternative quinol oxidase CydAB is used under microaerobic conditions. The regulation of synthesis occurs at the transcriptional level by the combined action of the oxygen sensors ArcBA and FNR (44). The catalytic subunits CyoAB contain the redox groups (heme a, heme o3, CuB) required for the intramolecular electron transfer, reduction of O2, and H+ pumping. The small accessory subunits CyoC and CyoD are not involved in catalysis. CyoABCD has (similar to the oxidase CydAB) no extended peripheral regions, and the major part of the enzyme is membrane integral (Fig. 5A to C and Fig. 5D). O2 is a lipophilic substrate and reacts at a site embedded within the membrane, and therefore a hydrophilic head typical for enzymes reacting with hydrophilic substrates is unnecessary. Function and structure of the enzyme are described in detail in chapter Respiration of Nitrate and Nitrite.
Nitrate reductase NarGHI and quinol oxidase CydAB function as redox-loop enzymes, and for the alternative (aerobic) nitrate reductase NarZYV, redox-loop function can be presumed (Fig. 5A to C and Fig. 5D). Nitrate reductase NarGHI has been studied biochemically, genetically, and structurally. The narGHJI operon encodes the structural proteins and the chaperone NarJ for cofactor insertion, assembly, and membrane insertion of NarGHI (131, 139). Synthesis of NarGHI is induced by the presence of nitrate under anaerobic conditions by the NarXL (and NarPQ) nitrate-responsive two-component systems (for a recent overview see, reference 200). Anaerobic expression is activated by the oxygen sensor FNR. NarGHI has low affinity for nitrate and is used under anaerobic conditions and high nitrate concentrations and is then the major enzyme during nitrate respiration (24). The structure of the enzyme is in many aspects a mirror image of FdnGHI with respect to composition, topology, intramolecular electron transfer, and proton potential generation (15, 18, 109, 110). The membrane-intrinsic subunit NarI consists of five TM helices (I to V) and anchors the enzyme to the membrane. The cytoplasmic subunits NarGH form the head of the enzyme extending into the cytoplasm. NarGH interacts with NarI by a hydrophobic interface and forms a stable heterotrimer. In contrast to FdnH, NarH contains no TM helix, and NarGH is fixed to the membrane only by hydrophobic interaction with NarI. The active sites for the quinol and nitrate are linked by eight redox centers forming a linear redox pathway of ~75 Å in length starting with heme bD (distal) at the quinol site, followed by heme bP (proximal), one [3Fe-4S] cluster, four [4Fe-4S] clusters, and the molybdenum MGD cofactor at the nitrate site. The center-to-center distances are generally ≤11 Å allowing rapid electron transfer (164). NarGHI forms (like FrdABCD) a "butterfly"-shaped dimer of the trimer [(NarGHI)2]. The redox centers of the two dimers are at least 33 Å apart, which precludes electron transfer between the dimers.
The Fe atoms of the heme b groups in NarI are coordinated by two His residues similar to FdnI, HydCWs, HyaC, and HybC. The ligands for the heme b molecules are located in TM II and V for each pair. The His pairs binding heme bD and heme bP are close to the distal (periplasmic) and proximal (cytoplasmic) side of the membrane, respectively. The essential quinol binding site is close to heme bD and the periplasmic face, but there appears to be a second quinol binding site (17, 174, 238, 239). The transmembrane arrangement of two heme B groups in the membrane anchor of NarI is typical for NarGHI-type enzymes and redox-loop subunits (15).
The overall reaction (QH2 + NO3− + 2H+in → Q + NO2− + H2O + 2H+ex) consumes two protons in the cytoplasm and releases two protons in the periplasm by a redox half-loop mechanism (2H+/2e−). Overall, the electron transfer chain from formate to nitrate generates 4H+/2e− by the combination of two redox loops of FdnGHI and NarGHI.
Nitrate reductase Z (NarZYV, narZYWV operon) is often referred to as the aerobic nitrate reductase. The operon encodes, in addition to the nar structural genes, the NarZ-specific chaperon NarW (20). The NarZYV proteins are highly similar to the NarGHI enzyme. NarZYV is a minor enzyme with low activity, and fully induced NarGHI is responsible for most (98%) of nitrate reductase activity (24, 95). NarZYV was not studied in detail, but it is assumed that the enzyme reacts similar to NarGHI. In the membrane anchor NarV the four His residues that bind heme b in NarGI are conserved, suggesting that the enzyme functions as a redox-loop enzyme as well. During exponential growth narZYVW is expressed at low levels under aerobic and anaerobic conditions and is not induced by nitrate. The enzyme therefore is regarded as a constitutive "aerobic" nitrate reductase. In stationary growth phase the enzyme is induced tenfold in a σS-dependent manner. The physiological role of the enzyme is not clear, but the studies suggest that NarZYV is used under general stress conditions and supports adaptation and survival by the use of nitrate respiration (33, 195).
Like many other bacteria, E. coli contains multiple terminal oxidases that are required for growth under different metabolic and physiological conditions (121, 153, 154). Quinol oxidase CyoABCD has relatively low affinity for O2 (KM = 0.2 μM) and is expressed at high pO2 (71, 98). At low pO2 or microaerobic conditions, quinol oxidase bd3 or CydAB, which has high affinity for O2, is induced (KM = 0.008 μM). CydAB is encoded by the cydAB genes. The oxidase is composed of the two catalytic subunits A and B comprising 17 or 18 transmembrane helices and small, soluble loop regions. CydAB is also in the content of prosthetic groups much simpler than CyoABCD and contains only heme b and heme d3 (121, 154). The enzyme is not a heme-Cu oxidase, and the redox reaction is therefore not coupled to proton translocation by proton pumping. It generates a proton potential by an orientation (or redox loop) mechanism by the release of 2H+ at the periplasmic face of the membrane during oxidation of the quinol, and consumption of 2H+during H2O formation by O2 reduction (153, 212). A homologous enzyme, AppBC encoded by the appBC genes has all structural and functional characteristic of CydAB and contains heme bd3. Upon (artificial) induction or overexpression, the AppBC protein can replace CydAB in microaerobic respiration and is functionally equivalent, but its physiological role is not clear (48, 205).
Most of the terminal reductases of E. coli do not conserve the free energy of the redox reaction in a proton potential. The enzymes have either cytoplasmic or periplasmic orientation (Fig. 5A to C and Fig. 5D).
Cytoplasmic orientation.
Fumarate reductase (FrdABCD) is described in detail in chapter Succinate as Donor; Fumarate as Acceptor. FrdABCD reduces fumarate, which is derived from glucose via PEP and oxaloacetate (32, 41, 87, 217). The end product succinate is one of the products of mixed acid fermentation. When external fumarate is used as an acceptor for respiration, an electron donor like H2, glycerol, or glucose is required in addition. Proteus rettgeri, however, dismutates fumarate to succinate and CO2 and is able to grow with fumarate as the only substrate (125). In fumarate respiration generation of a proton potential depends on the function of the dehydrogenases, like hydrogenase 2 (HybCOAB) or NADH dehydrogenase I. Succinate is not further metabolized under anaerobic conditions and excreted. The secondary carrier DcuB catalyzes an electroneutral antiport of fumarate against succinate when external fumarate is used (58, 102, 193). DcuC, on the other hand, excretes succinate2− in mixed acid fermentation in symport with protons (240).
Fumarate reductase is encoded by the frdABCD operon (see references in 41). The catalytic dimer FrdAB is anchored to the membrane by the hydrophobic proteins FrdCD. FrdAB is very similar in sequence, composition, and function to SdhAB (41, 49). Both dimers contain the same redox groups (FAD at the fumarate/succinate site, followed by one [2Fe-2S], [4Fe-4S], and [3Fe-4S] cluster each), which constitute the electron pathway from the fumarate/succinate sites to the membrane subunits (32, 99). The pathway is used in reversed direction in both enzymes.
The hydrophobic FrdCD proteins consist of six transmembrane helices that are contributed to the same part by FrdC and FrdD (32, 99). FrdDC differs significantly from SdhCD and contains no heme b. FrdCD binds, however, a quinone close to the cytoplasmic surface and the [3Fe-4S] cluster of FrdB. The heme b of SdhCD and the quinol site of FrdCD are at homologous positions and represent the active site for the reaction with the free substrate quinol (or quinone) from the membrane. The topology of the site in FrdCD results in the release of the protons from the quinol at the cytoplasmic side of the membrane without involvement of a redox loop. In FrdCD under some condition a second (distal) quinone binding site close to the periplasmic face of the membrane was identified, but only the proximal quinol site is regarded as functional. Overall, FrdABCD functions as an electron sink without energy conservation.
Periplasmic orientation.
Some terminal reductases are membrane bound with the catalytic site of the electron acceptor facing the periplasm. DMSO is used as an electron acceptor in combination with nonfermentatble cosubstrates as the electron donors. DMSO reductase (DmsABC) is a heterotrimer in which the catalytic dimer DmsAB is anchored to the membrane by DmsC. The enzyme is different from the DMSO reductase of the DmsAB CymA type of Rhodobacter species where the catalytic dimer DmsAB is in the periplasmic and interacts reversibly with the membrane-bound tetraheme C CymA (145, 146). Biochemical studies suggested a cytoplasmic location of DmsAB from E. coli, but later on their location on the periplasmic face of the membrane was shown (14, 175, 197). DmsC is membrane integral with eight transmembrane helices (227) and carries the site for the oxidation of the menaquinol (65, 177, 239). DmsA harbors a molybdenum cofactor at the active site for DMSO reduction. The DmsB protein closely interacts with the quinol oxidation site of DmsC and carries four [4Fe-4S] clusters that are assumed to transfer the electrons to DmsA to DmsC. The reaction of DmsABC presumably generates no proton potential similar to FrdABCD (Fig. 5A to C and Fig. 5D). YnfFGH, encoded by the ynfEFGHI operon, is a paralog of DmsABC (141). Bacteria overexpressing the ynf genes are able to complement a deletion of the dmsABC genes, and the overexpression strain is able to grow on DMSO as the electron acceptor, suggesting that YnfFGH is able to function as a DMSO reductase under specific conditions. The physiological conditions for YnfFGH synthesis and the function of YnfFGH have not been identified.
The catalytic subunits of nitrite reductase Nrf, nitrate reductase Nap and TMAO reductase TorA are located in the periplasm and not permanently linked to their membrane anchor or electron-donating membrane proteins (Fig. 5D). The enzymes are isolated as soluble proteins devoid of their membrane part which contains the quinone site and links the enzyme to the respiratory chain.
Trimethylamine N-oxide (TMAO) stems from marine animals and is used as an electron acceptor by the Tor respiratory system. The TMAO reductase TorCA has high specificity for TMAO and is encoded by the torCAD operon (150). TorA (TMAO reductase) is a periplasmic soluble enzyme with a molybdenum cofactor at the catalytic site. Folding and assembly of TorA occurs in the cytoplasm and requires the Tor-specific chaperone TorD (66, 92). TorC is a pentaheme C protein. It is fixed to the membrane by a N-terminal hydrophobic domain and is accessible to TorA from the periplasm. TorC interacts reversibly with TorA and transfers electrons from the quinones to TorA (69). TMAO reductase TorCA catalyzes TMAO reduction without formation of a proton potential. TMAO reduction is regarded as an anaerobic respiratory system, but the transcriptional regulation of the torCAD genes by O2 is not clear. Neither FNR nor ArcBA, which control the expression of other respiratory genes in response to O2, participate in the transcriptional regulation of the torCAD genes (166). The expression of the genes is transcriptionally activated by TMAO by using the TorSR two-component sensor systems together with the periplasmic-binding protein TorT (11, 112). TorYZ (torYZ genes) represents a second TMAO-specific respiratory terminal reductase (70). TorZ is a membrane-bound pentaheme C and a paralogue to TorC. The protein has been regarded previously as a minor biotin sulfoxide reductase. TorY (paralogue of TorA) has a periplasmic location. The torYZ gene cluster contains no paralogue of TorD, the specific chaperone of TorCA. TorYZ and TorCA are supposed to have a common ancestor. TorYZ is TMAO specific as TorCA, but both enzymes differ in some catalytic properties. The torYZ genes are expressed only at very low levels, and TMAO and DMSO do not induce the expression of the torYZ genes. Therefore, the physiological role of TorYZ is not clear.
The periplasmic nitrate and nitrite reductases NapABCGH and NrfABCD function under anaerobic conditions. Both enzymes have been suggested to couple the redox reaction to the formation of a proton potential in an unknown manner. Periplasmic nitrate reductase NapABCGH is the third nitrate reductase of E. coli (for review, see references 173 and 201). The Nap enzyme is composed of the soluble periplasmic NapAB with the nitrate site and a membranous part, NapCGH, which mediates electrons from the quinones to NapAB. NapABCGH has high affinity for nitrate and is expressed at low nitrate concentrations (170, 225). The enzyme has been suggested to represent the nitrate reductase when highly reduced C sources are metabolized in photosynthesis or in the presence of low nitrate concentrations. There are indications that the enzyme is able to conserve the redox energy in a proton potential (26). The Nap operon napFDAGHBC encodes seven proteins. NapAB represents the soluble periplasmic nitrate reductase. NapA contains the active site for nitrate reduction with a molybdenum cofactor and a [4Fe-4S] cluster, whereas NapB carries two heme C molecules. NapCGH constitutes the membrane integral part of the enzyme and provides the link to the quinols. NapF appears to be a cytoplasmic FeS-containing chaperonine that is required for FeS cluster assembly in the proteins before their export to the periplasm (158, 162). NapABCGH prefers menaquinol as the electron donor, whereas NarGHI favors ubiquinol. NapC is a tetraheme C protein and is suggested to transfer the electrons directly to NapAB. According to the model the menaquinol and ubiquinol sites are located separately on the NapCGH complex. NapC contains the active site for MKH2 and transfers the electrons directly to NapAB (27). NapGH contains the site for ubiquinol. NapG carries an FeS cluster and was suggested to transfer the electrons with NapH from the ubiquinol site to NapC, which then donates the electrons to NapAB (158, 162).
The sites for menaquinol and ubiquinol reduction are suggested close to the periplasmic side of the membrane, releasing the product protons to the periplasm without generating a proton potential (QH2 + nitrate → Q + nitrite + H2O). There are indications, however, for some proton-pumping activity by the enzyme (26, 27) which could depend on the NapGH subunits. Oxidation of MKH2 or UQH2 by nitrate would provide sufficient free energy for driving proton translocation of ≥1H+/1e− (compare Table 2).
Table 2Energetic parameters for the terminal reductases and oxidases of E. coli and S. enterica serovar Typhimurium |
E. coli is capable of nitrite ammonification similar to other γ-proteobacteria by using a respiratory nitrite reductase NrfABCD (see a comprehensive review in reference 191). NrfABCD is encoded by the nrfABCDEFG gene cluster (47, 91), which is expressed under anaerobic conditions in the presence of nitrite and nitrate. Under similar conditions a second nitrite-ammonifying enzyme is produced, the cytoplasmic nitrite reductase NirBD catalyzing NADH-nitrite reduction (40, 165, 224). NirB contains FAD, FeS clusters, and a siroheme as cofactors (100). The enzyme functions in detoxification of the nitrite produced by nitrate reductase NarGHI in the cytoplasm (40). NirBD has high activity and is responsible for most (>80 %) of the nitrite reductase activity (2, 168, 169).
NrfABCD nitrite reductase is composed of the membrane-bound NrfBCD complex and the periplasmic NrfA protein. NrfA is a pentaheme C, which is found preferentially in the soluble cell fraction (63, 114, 138). The protein is a homodimer with the five covalently bound heme C groups in close proximity (10, 55), enabling rapid electron transfer between the heme C groups. The electrons are donated by the NrfBCD complex to heme C2, and heme groups 3 to 5 transfer the electrons to the nitrite reduction site (heme C1). Heme groups 2 to 5 are axially ligated by two His residues each and function only in electron transfer. Heme C1 reduces nitrite and carries a nearby Ca2+ ion that is required for catalysis. The active site is accessible from the water space by a positively charged channel forming the nitrite pathway and a negatively charged channel by which ammonia leaves the active site.
The membrane-bound NrfBCD complex is supposed to function as a menaquinol-NrfA oxidoreductase. Most functional and structural details were derived from genetic information, and biochemical data are missing. According to the data NrfD is a membrane integral with eight transmembrane helices and represents the membrane anchor of NrfBCD. NrfB is a hydrophilic pentaheme C protein that is bound to the membrane by interaction with NrfD (78, 94). NrfC contains 16 conserved Cys residues and is supposed to function in electron transfer. The location and the binding of NrfB and C to the membrane are not known (91, 169). NrfB is suggested as the direct electron donor from NrfBCD to the periplasmic NrfA. NrfD could function as the quinol oxidase, and NrfC in the electron transfer between NrfD and NrfB. The protons from quinol oxidation might be released to the periplasmic side, rendering quinol-nitrite reduction electroneutral (QH2 + 1/3 HNO2 → Q + 1/3 NH3 + 2/3 H2O). It cannot be excluded that NrfD is a proton pump and uses the redox energy for generating a proton potential. The well-characterized nitrite reductase of W. succinogenes contains a NrfA very similar to the NrfA of E. coli (56, 191, 192). The membrane anchor and menaquinol site are formed by one subunit only, which does not function as a proton pump.
The respiratory enzymes of S. enterica have not been studied in much detail for the most part. The genomic sequence of S. enterica indicates the presence of all respiratory dehydrogenases from E. coli with the exception of malate:quinone reductase Mqo, TMAO reductase TorZY, and DMSO reductase YnfFGH for which the structural genes are missing. The respective alternative enzymes (NADH-dependent malate dehydrogenase, TorCA, and DmsABC) with well-characterized functions in E. coli are present in S. enterica. The physiological consequence of Mqo, TorZY, and YnfFGH deficiency in S. enterica is not known, since their function in E. coli is not clear as well. All other respiratory dehydrogenases appear to be present in a functional form as concluded from sequence and the predicted proteins. For the FHL, FHL-2, and Rsx complexes of E. coli, the corresponding gene clusters are also found in S. enterica. S. enterica is able also to use ethanolamine and 1,2-propandiol as (indirect) electron donors for respiratory growth, in particular, for tetrathionate respiration (171). Ethanolamine and 1,2-propandiol are fermented in a vitamin B12-dependent pathway to acetyl-CoA and propionyl-CoA, which are then oxidized in the citric acid/glyoxylate cycles and methylcitrate/citric acid cycles, respectively. The pathways provide NADH as the respiratory donor for anaerobic respiration and succinate.
A major difference is the use of tetrathionate and thiosulfate as electron acceptors for anaerobic respiration by S. enterica (Fig. 6) but not by E. coli. All other terminal reductases and oxidases encoded by E. coli are retained in S. enterica. Tetrathionate reduction and formation of sulfide is of diagnostic significance for differentiation of the closely related E. coli and S. enterica. Tetrathionate is reduced in three steps to sulfide (overall reaction: S4O62− + 18 [H] → 4HS− + 6H2O + 2H+). The first two steps are catalyzed by membrane-bound terminal reductases by using quinols as the electron donor. Tetrathionate reductase TtrABC (ttrBCA operon) reduces tetrathionate by two electrons to thiosulfate (S4O62− + QH2 → 2 HSSO3− + Q). Tetrathionate reductase requires anaerobic conditions for expression and has been studied by genetic means (85, 86). The sequence suggests that the catalytic subunit TtrA contains a MGD cofactor and a [4Fe-4S] cluster. TtrB comprises four Cys clusters typical for the binding of [4Fe-4S] clusters. TtrC is an integral membrane protein with nine predicted TM helices. TtrC contains no conserved His residues and no indications for the binding of heme b or other cofactors. It is suggested that TtrC contains the active site for quinol oxidation and serves as the anchor for the catalytic dimer TtrAB at the periplasmic face of the membrane. Topology, composition, and the function of the enzyme would be similar to DMSO reductase DmsABC. Overall, the reaction of the enzyme is supposed to be electroneutral without generation of a proton potential.
Thiosulfate is either supplied as the product of tetrathionate reduction, or stems from the medium. Thiosulfate reductase (PhsABC) converts thiosulfate to sulfide and sulfite (S2O32− + QH2 → HS− + HSO3− + Q), using quinol as the electron donor (38). PhsABC is a product of the phsABC gene cluster. PhsAB shows significant sequence similarity to the catalytic dimer DmsAB of E. coli DMSO reductase, suggesting that PhsB is a FeS-containing electron transfer protein and that the thiosulfate site with a molybdenum cofactor is located on PhsA (3, 83). PhsC has five transmembrane helices; it is highly hydrophobic and anchors PhsAB to the periplasmic side of the membrane. PhsC contains four conserved His residues that can be located in transmembrane helices 3 and 5. The supposed arrangement of the heme b groups is reminiscent of redox-loop enzymes like FdnGHI and NarGHI (see Fig. 8). The same arrangement is found also in SdhABC from MK-containing bacteria like Bacillus and anaerobic gram-negative bacteria like Desulfovibrio. SdhABC from these bacteria functions in a reversed redox-loop mechanism that consumes the proton potential to drive the endergonic reduction of MK by succinate (185, 237). The thiosulfate site of PhsABC is supposed to be in the periplasm (Fig. 6). Accordingly, the distal heme B for QH2 oxidation would be close to the cytoplasmic side of the membrane. The protons from QH2 oxidation would be released to the cytoplasm, and the substrate protons for thiosulfate reduction are consumed in the periplasm. In this way the reaction of PhsABC becomes electrogenic and is driven by the degradation of the proton potential. The overall reaction of thiosulfate reductase by MKH2 is highly endergonic under standard conditions (▵E0' MKH2/thiosulfate = 328 mV). By the reverse redox-loop mechanism the reaction would be supported by an input of (estimated) −170 mV from the proton potential, thereby decreasing the endergonic nature of the reaction significantly. It is suggested therefore that thiosulfate reductase uses a reverse redox-loop mechanism as shown in Fig. 6.
Anaerobic sulfite reductase (Asr) catalyzes sulfide formation, the final step of tetrathionate reduction [HSO3− + 3NAD(P)H → HS− + 3 NADP + 3H2O]. The enzyme is cytoplasmic and not linked to the respiratory chain (82, 85).
E. coli and Salmonella synthesize three different quinones (Q) (147, 214, 221), namely the benzoquinone ubiquinone (UQ) and the naphthoquinones menaquinone (MK) and demethylmenaquinone (DMK). The major part of the quinones contains a side chain of eight isoprene units (UQ-8, MK-8, and DMK-8) (43). The quinones serve as the redox mediators between the dehydrogenases and terminal reductases. The chemistry for the redox reaction (Q + 2e− + 2H+ ↔ QH2) is the same for all quinones. The respiratory enzymes donate or accept only the electrons and the protons are derived from or released to the surrounding. The quinones differ in their midpoint potentials (E0'), which range from +113 mV for the pair UQ/UQH2, to −74 mV for MK/MKH2. The midpoint potential of DMK is intermediate (+36 mV). The redox potential restricts reaction partners for the quinones: e.g., only the electronegative MKH2 (and DMKH2) are appropriate electron donors for fumarate respiration. The midpoint potential of fumarate (E0' = +30 mV) is too electronegative for efficient reaction with UQH2. On the other hand, succinate is too electronegative for substantial reduction of MK. Therefore, succinate dehydrogenase uses UQ as the preferred acceptor.
The quinone sites are at the membrane integral parts of the respiratory enzymes, or at the membrane/protein interface. The quinone sites of the membrane integral enzymes are either at heme b groups (e.g., in FdnI, NarI, or SdhCD), close to an FeS cluster with a nearby bound quinone (FrdCD), or not identified in detail. In one-subunit enzymes the active site is normally at the interface between the membrane and the dehydrogenase close to FAD, FMN, or PQQ cofactors, or to Cys thiols, which are also accessible from the water space by the hydrophilic substrates (compare Fig. 2, Fig. 3, and corresponding text).
The essential quinone in aerobic growth of E. coli is UQ (about 0.4 μmol/g dry weight), followed by DMK (0.2 μmol/g dry weight), whereas the content of MK is very low (<0.1 μmol/g dry weight) (214). The quinone contents are affected by the type of electron acceptor present in the medium, the growth phase, and the available C source. During fumarate or DMSO respiration MK (0.7 μmol/g dry weight) is the major quinone, followed by DMK (0.2 μmol/g dry weight), whereas the UQ contents are low. In nitrate respiration all quinones are found in high concentrations with DMK (0.7 μmol/g dry weight) as the major species.
The terminal reductases show a clear preference for specific quinones (Fig. 7), which was identified by the use of mutants that are devoid of specific quinones (80, 129, 148, 213, 221, 229, 230). The response can be explained mostly by the redox potentials of the quinones and of the acceptors. In aerobic growth UQ is the most efficient quinone for aerobic respiration, but the naphthoquinones support growth and electron transport to some extent (221). In anaerobic respiration with fumarate, DMSO, TMAO, nitrite, thiosulfate, and presumably tetrathionate, the naphthoquinones (MK + DMK) are required as the respiratory quinones and cannot be replaced by UQ to significant extent (80, 129, 147, 148, 161, 213, 229). Fumarate and TMAO reductases are able to use MK and DMK as well, whereas DMSO reductase prefers MK (230). Nitrate respiration with NarGHI as the major enzyme functions with UQ and MK, but for unknown reasons, not with DMK, which is intermediate in redox potential (221, 230). The periplasmic nitrate reductase NapABCDGH, on the other hand, prefers naphthoquinols as the electron donors compared with UQ (26).
The respiratory enzymes are combined in the membrane in many different combinations resulting in electron transfer chains with H+/2e− ratios varying from 2H+/2e− to 8H+/2e− in theory. Combination of NuoA-N with oxidase CyoABCD results in an overall ratio of 8H+/2e−, which is close to the ratio of 10H+/2e− from mitochondrial cytochrome c oxidase pathways. When NADH dehydrogenase II, the isoenzyme of NuoA-N is used, the H+/2e− ratio decreases from 8 to 4H+/2e− for aerobic respiration with oxidase CyoABCD. The same ratio of 4H+/2e− is achieved when CyoABCD is combined with a noncoupling dehydrogenase like Gcd, GlpD, SdhABCD, and others from the many noncoupling dehydrogenases. Under microaerobic conditions, when CydAB is active instead of CyoABCD, the H+/2e− ratios generally are decreased by 2H+/2e−.
Redox half-loop enzymes play an important role in proton potential formation in the respiratory chain of E. coli, in particular, under anaerobic conditions. Electron transfer chains composed of two redox-loop enzymes, e.g., FdnGHI and NarGHI in formate-nitrate respiration (Fig. 8) yield 4H+/2e−. Combination of a redox-loop enzyme with a noncoupling enzyme, e.g., FdnGHI with FrdABCD, DmsABC, or NapABCGH (assuming that NapABCGH is a noncoupling enzyme), or the combination of SdhABCD with CydAB, always yields 2H+/2e−, irrespective of the orientation and topology of the noncoupling enzyme (Fig. 9).
On the other hand, combination of a nonelectrogenic (or noncoupling) dehydrogenase with a nonelectrogenic terminal reductase never results in proton potential formation, irrespective of the orientation of the enzymes (Fig. 10). Thus, NADH-fumarate and NADH-DMSO respiration by a (hypothetical) combination of Ndh with FrdABCD or DmsABC results in an H+/e− ratio of 0 for both combinations despite the inverse orientation of FrdABCD and DmsABC. These combinations are not found under physiological conditions since the enzymes are not produced at the same time. Similarly, for the combination of the periplasmic glucose:quinone reductase Gcd with noncoupling terminal reductases like FrdABCD or DmsABC, no formation of a proton potential is predicted. Therefore, combination of noncoupling dehydrogenases with noncoupling terminal reductases is never effective in proton potential formation, and proton potential formation by a mere topology-based mechanism of separate enzymes is not verified. Combination of suitable enzymes according to these criteria apparently is guaranteed by transcriptional regulation, whereas the combination of unsuitable enzymes is avoided (216).
Overall it appears that in aerobic respiration H+/2e− ratios of 4 to 6 are common for many respiratory chains, whereas in anaerobic respiration ratios of 2 to 4 are frequently found. This means that the H+/2e− ratios of the aerobic quinoloxidase pathway or respiratory chain of E. coli are often only half that of the mitochondrial respiratory chain. The extremes of H+/2e− ratios of the E. coli respiratory chain (8H+/2e− and 0H+/2e−) are avoided by transcriptional regulation, which generally prevents synthesis of the corresponding enzyme combinations.
Growth yields, H+/e− and ATP/e− ratios, and other energetic parameters have been studied for a long time and in great detail, in particular, for aerobic, nitrate, and fumarate respiration of E. coli. The studies are complicated by the large number of electron transport systems expressed at the same time and the presence of isoenzymes with different H+/e− ratios. Previous reviews summarize energetics and the H+/e− and ATP/e− ratios for various respiratory systems (68, 216). Here, we concentrate on information derived from topology of the enzymes, the substrate sites, and the topology of the H+-releasing and -consuming reactions, which, in combination with biochemical, structural, and bioinformatic data, give reliable information on the theoretical and maximal H+/2e− ratios. It has to be considered, however, that the actual values are often smaller due to H+ leakage or losses on other sites.
Free energy of aerobic respiration is much higher (−2,870 kJ/mol for glucose oxidation to CO2) than for anaerobic respiration or mixed acid fermentation (1 glucose → 1 acetate + 1 ethanol + 2 formate, ▵G0' = −218 kJ/mol of glucose). Nevertheless, the energetic parameters (proton potential, ▵p, and phosphorylation potential, ▵GP') of E. coli cells are similar for aerobic and anaerobic respiration. ▵GP' for respiration with O2, nitrate, fumarate, DMSO, and TMAO is the same within a small range from 46.2 to 47.7 kJ/mol when determined under comparable conditions from the cellular ADP, ATP, and Pi contents (39, 84, 211). The proton potential produced by aerobic and anaerobic respiration varied only slightly; in aerobic respiration, values around −0.17 V were measured, which decreased by only 0.1 to 0.2 V in anaerobic respiration. The theoretical or maximum H+/e− ratio [or (nH+/ne)max] for H+ translocation by respiratory enzymes can be calculated according to the equation (1): (nH+/ne)max = ▵E0'/▵p. Assuming that proton potential is generally about −0.17 V under respiratory conditions, H+/e− ratios of 1 or 2 require a minimum ▵E0' of the redox reaction of 0.17 V and 0.34 V, respectively, under equilibrium conditions.
FoF1 ATP synthetase of E. coli is composed of 10 to 11 subunits C in the C ring of the membrane rotor, each transducing 1 H+ per turn (106, 149). One complete rotation of the enzyme is coupled to the formation of 3 ATP, resulting in a H+/ATP stoichiometry of 10 to 11 H+/3 ATP, or about 3.5 H+/ATP. The value is close to experimental values (118). By combining the H+/e− ratio with the H+/ATP ratio of 3.5 (i.e., by dividing the H+/e− by 3.5), the (ATP/e−)max ratio can be calculated for each reaction or respiratory chain. The experimental ATP/e− ratios are often significantly lower. We will deal here with the theoretical or maximal H+/2e− and ATP/2e− ratios since other physiological parameters and reactions causing a decrease in the ratio are not subject of the review.
Tables 1 and 2 show the midpoint potentials of the donors and acceptors of the respiratory chains of E. coli and S. enterica serovar Typhimurium and some of the resulting energetic parameters. The electron donors cover a wide range from −432 mV to +30 mV (Table 1). Considering the ▵E0' for reaction with UQ and MK, it becomes obvious that UQ is suited for accepting electrons from all donors, whereas MK is too electronegative for efficient reaction with succinate. This estimation is supported by the requirement for UQ for growth on succinate (221, 229). The ▵E0' values (donor–UQ) demonstrate that a considerable number of redox reactions do not supply sufficient ▵E0', or free energy, for driving H+ translocation, since ▵E0' is below 170 mV (see above). This applies to the oxidation of succinate with UQ as the acceptor, and to the oxidation of glycerol-3-P, lactate, malate, glucose, and alanine by MK. Only the electronegative donors with E0' ≤ −320 mV (formate, H2, and NADH) are used for conserving the redox energy by proton potential-forming enzymes. Remarkably, the most electronegative donor pyruvate is oxidized by a non-energy-conserving enzyme (PoxB). NADH, which is at the lower limit of donors that are used for energy conservation, is oxidized by a coupling (NuoA-N) or a noncoupling (Ndh) enzyme. Glycerol-3-P, lactate, malate, and glucose have ▵E0' values with UQ as the acceptor, which are sufficient to drive H+ translocation according to the above arguments (▵E0' > 170 mV). In vivo, however, the difference is apparently not sufficient to exploit the redox energy for proton potential formation. Pyruvate, which is a very negative electron donor, does also not use the high ▵E0' for conservation of the redox energy.
The electron acceptors show also a large range in redox potential as well, ranging from +24 mV (tetrathionate) to +820 mV for O2 (Table 2). The midpoint potential of the pair thiosulfate/sulfite + sulfide is very negative and represents a specific situation. The electronegative thiosulfate can be used as an acceptor only after activation of the electrons, presumably by a reversed redox-loop mechanism (see Fig. 6). The reaction of UQH2 with tetrathionate and fumarate would be endergonic under standard conditions, and with TMAO and DMSO only weakly exergonic. With MKH2 as the donor, the reactions are all exergonic (apart from thiosulfate as described above) and MKH2 is for energetic reasons the preferred electron donor for less electro-positive acceptors. This argument is in agreement with the growth deficiency of men mutants (MK deficient) on tetrathionate, fumarate, DMSO, and TMAO. Nitrite reduction would be able to use MK and UQ as well from a thermodynamic point of view, but only MKH2 supports nitrite reduction by the NrfABCD enzyme (213).
Assuming that ▵E0' of a redox reaction has to be 200 mV or higher for coupling the free energy in a proton potential, then thiosulfate, tetrathionate, fumarate, TMAO, and presumably also DMSO reduction by the quinols would not be coupled to proton potential formation. This is in agreement with the experimental results. The oxidases CyoABCD and CydAB and nitrate reductase NarGHI using electropositive acceptors, however, are coupling enzymes. The redox energy of nitrite reduction by NrfABCD would be sufficient for coupling the reaction to the formation of a proton potential. In particular, with MKH2 the ▵E0' value would suffice for energy conservation.
In summary, ▵E0' of a redox reaction has to be significantly higher than 200 mV in individual enzyme reactions for conservation of the redox energy in a proton potential. Accordingly, most of the dehydrogenases and terminal reductases of the respiratory chain do not use the redox energy for proton potential formation. The corresponding dehydrogenases contribute to the formation of a proton potential only when combined with energy-conserving terminal reductases like NarGHI and the oxidases. Accordingly, low-potential donors are used preferentially in aerobic or nitrate respiration. The same applies to the use of noncoupling terminal reductases, like fumarate, TMAO, and DMSO reductase, in combination with coupling dehydrogenases. Thus, in fumarate respiration only NuoA-N is used, whereas in aerobic respiration coupling (NuoA-N) and noncoupling (Ndh) NADH dehydrogenases operate (73, 210).
In principle, any of the dehydrogenases can be coupled to any of the terminal reductases, provided that the enzymes are expressed under the same conditions and able to use the same type of quinone. Thus, succinate dehydrogenase can be linked only to terminal reductases that accept UQ as the redox mediator. By the variation of the enzymes the H+/e− ratios can be varied to a considerable extent to adjust the cellular energetics (Table 3). By combination of NADH dehydrogenase I (NuoA-N, 4H+/2e−) with oxidase CyoABCD (4H+/2e−), an overall yield of 8H+/e− can be achieved, which is the highest theoretical value for the (aerobic) respiratory chain of E. coli. However, in aerobic respiration the noncoupling NADH dehydrogenase II has high activity, resulting in an overall yield in the range from 4 to 8 H+/2e−.
Table 3Some typical respiratory chains of E. coli: composition from enzymes and resulting H+/2e− ratios |
Linking of noncoupling dehydrogenases like Ndh to noncoupling terminal reductases like fumarate reductase (overall H+/e− = 0) is avoided as well. During fumarate respiration only NuoA-N is produced, enabling a H+/2e− ratio of 2 for NADH-fumarate respiration. In these pathways, exergonic but noncoupling enzymes have a positive effect on the energetics by their high-flux rates and their effect on the steady state of respiration and the energetics of the other coupling reactions of the respiratory chain.
The energetic considerations are verified by the corresponding transcriptional regulation. Noncoupling terminal reductases like fumarate or DMSO reductase are produced preferentially in combination with energy-conserving dehydrogenases like FdnGHI, hydrogenase 2 HybCOAB, and NADH dehydrogenase I. Vice versa, the energy-conserving oxidases are produced often in combination with noncoupling dehydrogenases for low-potential substrates.
Following are some useful databases:
http://www.ecocyc.org/; http://ecogene.org/; http://cmr.tigr.org; http://beta.uniprot.org; http://genolist.pasteur.fr/Colibri/; http://www.uni-giessen.de/~gx1052/ECDC/ecdc.htm; http://www.jgi.doe.gov/; http://www.ncbi.nlm.nih.gov/; http://www.sanger.ac.uk/; http://www.expasy.org/sprot/; http://ecoli.aist-nara.ac.jp/; http://cmr.tigr.org/tigr-scripts/CMR/GenomePage.cgi?database=ntec01; http://swift.cmbi.kun.nl/swift/genequiz/; http://gib.genes.nig.ac.jp/single/index.php?spid=Ecol_K12_MG1655; and http://xbase.bham.ac.uk/
We are grateful to S. Lux (Mainz) for help in preparing the figures and to D. Jahn (Braunschweig) for helpful discussions.
Work in the authors’ laboratory was supported by grants from Deutsche Forschungsgemeinschaft.
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Module3.2.2
