Fermentation
Chapter
18
AUGUST BÖCK and GARY SAWERS
Fermentation was orginally described by Pasteur as "la vie sans air." It is a condition under which growth occurs without any exogenous electron acceptor. Therefore, certain metabolic constraints are necessarily placed upon the organism that flourishes by this mode of growth. The enterobacteria with their remarkably flexible metabolism are, together with the clostridia, the classical organisms of bacterial fermentation. This chapter will concentrate on the fermentation principally of Escherichia coli and Salmonella typhimurium. Since studies have revealed that they are essentially indistinguishable with regard to their fermentative metabolism, the chapter has been written with this in mind. Other genera of the enterobacteria, however, have evolved specialized fermentative pathways, not common to either E. coli or S. typhimurium. Thus, Klebsiella and Citrobacter species are capable of fermenting glycerol, while Enterobacter and Klebsiella spp. produce the alcohol 2,3-butanediol. Therefore, we have also included sections which discuss these pathways in the light of energy generation and the maintainance of redox balance.
The oxidation of hexoses, such as glucose, to pyruvate by the Embden-Meyerhof-Parnas pathway generates two molecules of NADH (Fig. 1). To maintain glycolytic flux, the oxidized form of this pyridine nucleotide must be regenerated, and during fermentation, this is achieved by depositing the reducing equivalents on partially oxidized metabolic intermediates, which are then excreted from the cell. The fermentation products of E. coli comprise a mixture of ethanol and acetic, formic (H2 + CO2), lactic, and succinic acids (34, 174). These products have different oxidation states (Table 1), and by adjusting the proportion of each compound produced, E. coli can modulate its metabolism to grow fermentatively on a variety of compounds such as hexitols and hexonic and hexuronic acids.
Table 1Oxidation states of various substrates and products of E. coli fermentation |
The degradation of glucose generates two molecules of pyruvate and two molecules of NADH (effectively four reducing equivalents [H]), the latter arising as a consequence of the oxidation of glyceraldehyde 3-phosphate by glyceraldehyde-3-phosphate dehydrogenase (Fig. 1). The more reduced sugar alcohol, glucitol, enters the cell via the phosphoenolpyruvate:sugar phosphotransferase system (PTS) but then must be oxidized before it enters the Embden-Meyerhof-Parnas pathway as fructose 6-phosphate (see chapter 20, this volume). The dehydrogenation of glucitol 6-phosphate generates a further molecule of NADH over and above those produced during glycolysis (132, 210). Consequently, fermentation of glucitol requires that more reduced fermentation products be excreted to accommodate the recycling of NAD+. Fermentation of sugar derivatives more oxidized than glucose, for example gluconate or glucuronate (Table 1), has the advantage for the cell that less NADH is produced (3, 34, 174). Gluconate fermentation generates only one molecule of NADH, as does the fermentation of glucuronate; however, the metabolism of glucuronate requires that it be first reduced via a fructuronate intermediate to mannonate by d-mannonate dehydrogenase (79), thus consuming a molecule of NADH. The result is that conversion of glucuronate to two molecules of pyruvate is effectively redox balanced (Fig. 1). The advantage for the cell that the fermentation of these latter two compounds affords with regard to maintaining redox poise is counterbalanced by their limited energy yield (see below), since they must be metabolized via the Entner-Doudoroff pathway (Fig. 1). These examples show quite clearly that fermentation is a compromise between attempting to attain maximal energy yields at the cost of upholding redox balance.
The oxidative decarboxylation of pyruvate by the pyruvate dehydrogenase (PDH) complex generates one NADH and as a consequence is functionally restricted to respiratory metabolism (71, 95). The activity of the PDH complex is negatively regulated by NADH (73). Nonoxidative cleavage of pyruvate to acetyl coenzyme A (acetyl CoA) and formate by the pyruvate formate-lyase (PFL) enzyme, therefore, is of clear benefit to E. coli during fermentative growth (chapter 15; 101).
The synthesis of PFL is enhanced 10- to 15-fold by a shift to anaerobiosis, and this regulation is mediated at the transcriptional level (101, 159, 160). The transcription factors FNR and ArcA positively regulate transcription from the seven promoters that transcribe the pfl gene (157, 160, 162, 163). Anaerobic pyruvate accumulation further enhances transcription of pfl, and the integration host factor is required to mediate this process (173).
The PFL enzyme is functional only anaerobically (chapter 15; 101), and activity is strictly controlled by a remarkably sophisticated interconversion cycle. PFL must be activated by introduction of a stable organic free radical into the polypeptide chain (205). This reaction is catalyzed by an iron-dependent activating enzyme that requires reduced flavodoxin and S-adenosylmethionine as substrates. Removal of the radical and conversion of the enzyme to the inactive, oxygen-stable species is catalyzed by alcohol dehydrogenase (ADH) (98, 99). The redox status of the cell clearly has a key function in controlling which of the two enzymes, PDH or PFL, is active under a particular condition. The mutually exclusive nature of the enzyme activities can be demonstrated quite readily in that mutants that cannot synthesize a functional PDH complex are auxotrophic for acetate when grown aerobically but are prototrophs during fermentative growth (105). On the other hand, pfl mutants are prototrophic for aerobic growth but grow very poorly by fermentation if the culture medium is not supplemented with acetate (95, 124, 202). Interestingly, when the redox potential of the culture medium is poised at +430 mV during anaerobic growth with nitrate, which lies between the extremes found during aerobic (+820 mV) and fermentative (–420 mV) growth, then either of the two enzymes can be functional in metabolizing pyruvate (95).
To attain redox balance, all of the reducing equivalents must be accounted for in the products formed. E. coli has several possibilities available for reoxidizing NADH, and Fig. 1 illustrates this metabolic potential. For every molecule of pyruvate cleaved by PFL, one molecule of formate and one molecule of acetyl CoA are formed; therefore, a maximum of one third of the carbon from glucose can be converted to formic acid. Formate either is excreted from the cell (14, 189) or is decomposed to carbon dioxide and dihydrogen by the formate hydrogenlyase (FHL) complex (164, 180, 181).
Acetyl CoA has two alternative fates (Fig. 1). The energy in the thioester bond can be conserved in the form of ATP by the action of the phosphotransacetylase (PTA) - acetate kinase (ACK) pathway. Acetate is the final product of this pathway, but its formation does not result in consumption of any reducing equivalents. Alternatively, the energy can be sacrificed by reducing acetyl CoA to ethanol through the two dehydrogenation reactions catalyzed by ADH (Fig. 1). It is unlikely that acetaldehyde is released as an intermediate in vivo (but see reference 41), due to its potential toxicity. Therefore, although both reactions of this branch are reversible, their use commits the cell to consuming four reducing equivalents (two NADH). Ethanol, therefore, is the most highly reduced major fermentation product of E. coli.
The other possibilities for disposing of reducing equivalents are as lactate or succinate (Fig. 1). The lactate dehydrogenase (LDH) reaction results in the reoxidation of one NADH but, like the ADH reaction, has the disadvantage that it squanders an energy-rich pyruvate molecule. Succinate normally makes up only 5 to 10 mol% (Table 2) of the fermentation products, and the vast majority produced by fermenting cultures is derived from phosphoenolpyruvate (3, 34, 174). Use of this latter pathway exclusively would not permit growth by fermentation. The first step in this route is the carboxylation of phosphoenolpyruvate, with the large proportion of the CO2 arising from the FHL reaction. Hence, the availability of metabolic CO2 limits the amount of succinate production. The subsequent malate dehydrogenase and fumarate reductase reactions each consume two redox equivalents per succinate formed (Fig. 1). It is also theoretically possible to generate succinate by the condensation of two acetyl CoA units via the glyoxylate cycle. This is not a feasible alternative, however: first, it produces two reducing equivalents, and second, the glyoxylate cycle is strongly glucose repressed (121; chapter 21).
Table 2Calculation of the fermentation balance for growth of E. coli on glucose |
Examination of the oxidation states of the various substrates and products of fermentation shown in Table 1 gives a clear idea of the alternatives available to E. coli to balance its fermentation. For example, it should be possible to balance most fermentations simply by varying the proportions of acetate and ethanol produced, which would imply that the LDH reaction is superfluous. Indeed, Clark and his colleagues have demonstrated that mutants unable to produce the fermentative NADH-dependent LDH have no obvious anaerobic growth defects (124). This finding is also in accord with the regulation of LDH enzyme synthesis and activity, both being activated by a low pH (124). Hence, during growth of E. coli in batch culture lactate is the organic acid which appears subsequent to all of the others (189). The reasons why the LDH reaction has been conserved during evolution probably include (i) the reaction gives an enhanced metabolic flexibility to the organism, and (ii) perhaps more importantly, the production of lactate is of value energetically (see Energy balance, below).
Studies with ack, adh, and pta mutants have demonstrated the essential requirement of the PTA, ACK, and ADH enzyme reactions to permit growth of E. coli on certain fermentable sugars. Mutants unable to synthesize ADH cannot grow anaerobically on glucitol, glucose, or gluconate, but they can ferment the highly oxidized glucuronate (39, 72). This reflects the fact that metabolism of glucuronate to pyruvate is redox balanced (Fig. 1). Neither ack nor pta mutants can grow on any of the above substrates (36, 72). Strains defective in both pta and adh or adh and pfl recover the ability to ferment glucose (72, 137), and essentially such double mutants perform a lactate fermentation. It would be anticipated that the single pta or ack mutants would also grow on glucose by lactate fermentation simply through the bottleneck arising at PFL; clearly this does not occur. These findings indicate that if PFL is functional, the PTA-ACK pathway is essential to permit growth. This suggests that there may be some common allosteric regulatory mechanism controlling the activities of the PFL, PTA, ACK, and ADH enzymes. That communication between PFL and ADH occurs has been demonstrated recently (98, 99).
Determination of the fermentation balance provides information on the type of cleavage that occurs during the catabolism of a particular sugar by a particular organism. It has been determined that, to a close approximation, only 10% of the carbon source is assimilated due to the poor energy yields of fermentation. Therefore, fermentation balance studies are particularly useful in comparing and contrasting the metabolism of different sugars, or their derivatives, between different bacterial strains (35, 174).
The organism to be investigated should be placed in a sealed vessel under an inert atmosphere, and fermentation should be allowed to proceed to completion before the concentrations of the various products are determined. Gas-phase and high-pressure liquid chromatography procedures can be used to determine the concentrations of the compounds. Recently, in vivo nuclear magnetic resonance spectroscopy has been developed as a useful method for examining the progress of fermentation (3, 124, 133, 134).
An example of a fermentation balance determination for E. coli grown with glucose is presented in Table 2. The sum of the oxidized and reduced products (columns 4 and 5) should equal the oxidation state of the substrate. In the example presented the sum is +0.14, suggesting that not all products were efficiently recovered (Table 2). The carbon recovery is obtained by multiplying the moles of product determined by the number of carbon atoms in the product; in this case 91% of the carbon was recovered.
Analysis of the products shown in Table 2 indicates that succinate amounted to only 5% of the total product, while the acetate:ethanol:lactate:formate (+CO2) ratio was approximately 1:1:2:2 (Table 2). Hence, one third of the carbon from glucose was metabolized to lactate. The total moles of ethanol plus acetate equals the total moles of formate + CO2 produced, and this is in agreement with what would be anticipated from the cleavage of pyruvate by PFL (Fig. 1).
On the basis of theoretical considerations, fermentation with glucitol should yield an increased proportion of ethanol relative to acetate, since degradation of glucitol generates an extra NADH compared with fermentation of glucose (Fig. 1). Nuclear magnetic resonance studies indeed showed that the ratio of ethanol to acetate changed from 1:1 for growth with glucose to 6:1 for glucitol-dependent growth (3). The proportion of succinate also increased, and no lactate was produced relative to that found after growth with glucose. In sharp contrast, fermentation of the highly oxidized glucuronate resulted in an ethanol:acetate ratio of 1:5, with no succinate and only minor amounts of lactate being excreted (3). Metabolism of glucuronate to two molecules of pyruvate is redox balanced; therefore, there is no requirement to produce ethanol or lactate, and all of the acetyl CoA can be channeled through the PTA-ACK pathway to synthesize ATP (Fig. 1).
In growing cells, a large fraction of the metabolic energy is required for transport of metabolites and ion fluxes, as well as for biosynthetic purposes. Fermentation yields two ATP per glucose from glycolysis (Fig. 1). Acetyl CoA is therefore the primary "energy-rich" intermediate during fermentation (197), yielding potentially a further two ATP per glucose via the ACK reaction (Fig. 1). However, as discussed above, redox balance must be maintained, and as a consequence at least one molecule of acetyl CoA is sacrificed for the disposal of excess reducing equivalents. Thus, on average, the ACK reaction generates only one additional ATP per glucose. The fermentation of more reduced substrates necessitates a greater proportion of ethanol production with concomitantly less ATP generation via acetate kinase (Fig. 1). More oxidized substrates, on the other hand, such as gluconate or glucuronate, permit more ATP to be generated by substrate-level phosphorylation, but because these substrates are metabolized by the Entner-Doudoroff pathway only one ATP results from their degradation to pyruvate.
There is also a considerable energy potential stored in the reduced organic acids produced. Konings and colleagues have proposed that a substantial electrochemical proton gradient can be generated by carrier-mediated efflux of fermentation products, e.g., lactate, with protons (129). This energy recycling model is based on the assumption that the proton-solute-carrier complex is electroneutral and that the stoichiometry of proton to solute varies with the dissociation constant of the carrier. Thus, a carrier that dissociates at physiological pH could symport more protons than solute molecules, thereby creating an electrochemical proton gradient. Experimental validation of this proposal has been presented for l-lactate efflux employing E. coli membrane vesicles. The results are consistent with these processes supplying sufficient energy to meet the demands imposed by the transport of metabolites (196). Recent studies have identified a putative carrier for formate that is present in fermenting cells and that could function along the same principles (189).
LDH can be operationally classified into two types. NAD-dependent LDHs (nLDHs) are enzymes that catalyze the reduction of pyruvate to lactate, and there is no evidence to indicate that they catalyze the reverse reaction in vivo. In contrast, NAD-independent LDHs (iLDHs) catalyze the reverse reaction, i.e., the oxidation of lactate to pyruvate (61). Essentially, nLDHs are pyruvate reductases and iLDHs are dl-lactate oxidases.
E. coli has three LDH isoenzymes. Two isoenzymes are membrane-associated flavoproteins that couple the oxidation of dl-lactate to energy generation and consequently have a function in respiratory metabolism. One of these flavoproteins is specific for the l-(+)-lactate isomer, while the other specifically oxidizes d-(–)-lactate (60, 61, 131). The genes encoding both isoenzymes have been cloned and sequenced (29, 43).
The third isoenzyme is a d-(–)-nLDH (EC 1.1.1.28) that is soluble, couples pyruvate reduction to the oxidation of NADH, and has a purely fermentative function (193, 194, 195). Early studies indicated that d-(–)-lactate production by E. coli is enhanced at low pH (183, 198). More recently, Mat-Jan et al. (124) demonstrated that enzyme activity is strongly induced by anaerobiosis and a low pH in the culture medium. Acid induction of enzyme activity does not occur during aerobic growth, which may indicate that expression of the ldhA gene, encoding the enzyme, is enhanced during anaerobiosis (124).
d-(–)-nLDH has been purified and its kinetic properties have been characterized in some detail (193, 194, 195). The enzyme has an M r of 115,000 and is extremely sensitive to autoxidation. This may result from oxidation of the 12 thiol groups of the enzyme, all of which have been proposed to be exposed at the surface of the enzyme based on their reactivity toward alkylating agents (194). Both pyruvate and NAD afford the enzyme protection against alkylation by acetamide, indicating that the cysteinyl residues may be at or near the active site of the enzyme. Surprisingly, NADH enhances the rate of alkylation. Tarmy and Kaplan (194) have suggested that binding of the reduced coenzyme causes a conformational change in the protein.
Kinetic studies have shown that there are two sites on the enzyme that interact with pyruvate. One is the catalytic site, and the other is a site for allosteric control (195). α-Ketobutyrate is also capable of activating the enzyme, but oxamate, another pyruvate analog, cannot. Oxamate, however, is an inhibitor of catalytic activity with an apparent Km of 18 mM (195). The Km for pyruvate is 7 mM, which is very high (193, 194). The V max of the enzyme was found to decrease with increasing pH, suggesting that protons are also positive allosteric effectors of d-(–)-nLDH. The enzyme shows normal Michaelis-Menten kinetics toward NADH (195).
The complex kinetic features of d-(–)-nLDH are completely in accord with the physiology of fermentation (34, 133, 134). Only when the pH of the medium drops, due to increased concentrations of acetate, formate, and succinate, is the excretion of lactate initiated. The reduced flux of pyruvate through PFL, resulting in intracellular pyruvate accumulation, together with the acidic environment effects allosteric activation of the nLDH. The reduction of pyruvate to lactate is an important means of maintaining redox balance, since the reaction reoxidizes the NADH generated by the glyceraldehyde phosphate dehydrogenase step of glycolysis. Exactly this function of LDH was exploited to identify mutants deficient for the nLDH isoenzyme (124). Clark and colleagues mutagenized a strain that could not synthesize a functional PFL enzyme and screened for mutants no longer capable of growing anaerobically by fermentation (124). These mutants had drastically reduced or no d-(–)-nLDH activity. The mutation was localized to 30.3 min on the E. coli chromosome. By making use of this information, Clark and colleagues succeeded in cloning and sequencing the ldhA gene and showed that it encodes a polypeptide of 36,532 Da which exhibits significant homology with d-LDH from Lactobacillus bulgaricus and Lactobacillus plantarum (D. P. Clark, personal communication).
In anaerobically growing E. coli and S. typhimurium, acetyl CoA can be metabolized by one of two pathways (Fig. 1). One pathway involves the two-step reduction to acetate by ADH and will be discussed below. The second pathway comprises the enzymes PTA (phosphotransacetylase; acetyl CoA:orthophosphate acetyltransferase) and ACK (acetate kinase; ATP:acetate phosphotransferase) and is one of the major routes used by fermenting organisms to generate ATP (197). The pathway is reversible, enabling enterobacteria to grow with acetate as sole carbon source aerobically (26, 155). Enzyme synthesis is essentially constitutive, and it is likely, based on results obtained from mutant studies, that the genes encoding both enzymes form an operon that is located at 49.5 min on the chromosome of both E. coli and S. typhimurium (26, 70, 104, 111, 125, 137).
ACK.
ACK (EC 2.7.2.1) has been purified to homogeneity from both E. coli and S. typhimurium, and the properties of the enzyme from both sources are very similar (57). The molecular weight of 40,000 determined for the homogeneous enzyme is in good agreement with that (43, 297) deduced from the translation of the sequenced gene (125). The enzyme is probably functional as a homodimer that is cold labile (57). Studies performed with the inactivated enzyme demonstrated that this lability is dependent on the protein concentration and reactivation could be accomplished in the presence of MgATP and acetate (57). Similar behavior is exhibited by enzyme I of the PTS (103).
ACK has quite a high Km for acetate of 7 mM. In contrast, the Km for acetyl phosphate is 0.16 mM (57), which is in accord with the main function of the enzyme in vivo being ATP generation. The V max of the purified enzyme is similar (2,000 to 2,600 mmol–1 mg–1 of protein) in both the forward and reverse directions. The enzyme has a requirement for Mg2+ ions but is inhibited by Na+ and Li+. Mn2+ ions can substitute for Mg2+ effectively (57).
The results of several studies point to a phosphoenzyme intermediate participating in the reaction (4, 56, 88). It was demonstrated that both substrates had to be added to the enzyme before either product was released, which would be consistent with such an intermediate and suggests a sequential reaction mechanism (88). Studies performed with buffers of different pH indicated that the phosphate group is linked either to an aspartyl or to a glutamyl residue (56). Fox et al. showed that phospho-ACK could transfer the phosphate group to enzyme I of the PTS (56). This finding led the authors to speculate that sugar transport may be regulated by the energy charge (ATP/ADP ratio) and Krebs cycle intermediates (56). More recently, Wanner and Wilmes-Riesenberg have proposed that the activity of the PTA-ACK pathway may control the phosphate regulon and that acetyl phosphate might be an effector of gene regulation (207).
PTA.
PTA (EC 2.3.1.8) activity is extremely labile, but by using ammonium sulfate as a stabilizer it could be purified 610-fold from aerobically grown E. coli cells (171). The relative molecular size of the enzyme lies between M r 160,000 and 250,000 (171). Little is known about the subunit structure of PTA, but on the basis of genetic studies it is likely that it consists of a single type of polypeptide (26, 111).
The Kms of the enzyme for acetyl phosphate and CoA are 3 mM and 0.3 mM, respectively (171). Interestingly, it was found that the Km for acetyl phosphate is pH dependent, decreasing by one order of magnitude as the pH is reduced from 7.8 to 5.8 (190). Pyruvate is a positive allosteric effector, and NADH, ADP, and ATP are negative allosteric effectors of the enzyme. Pyruvate was found to lower the Km for acetyl phosphate while the V max remained unaffected (190).
PTA and ACK in Other Enterobacteria.
A study involving isolation of pta and ack mutants by Brown et al. indicated that the organization of the genes encoding pta and ack in Enterobacter aerogenes may be similar to that in E. coli and S. typhimurium (26, 27). A major difference between the organisms, however, is that unlike E. coli and S. typhimurium, pta ack double mutants of Enterobacter aerogenes cannot grow aerobically with acetate. This indicates that it lacks the enzyme acetyl CoA synthetase, which both E. coli and S. typhimurium possess (155). These studies with the pta ack mutants of Enterobacter aerogenes confirmed the importance of the amphibolic nature of the PTA-ACK pathway for enterobacteria and provided strong evidence to indicate that the synthesis of the butanediol pathway in Enterobacter aerogenes is strongly controlled by acetic acid (27).
The reduction of acetyl CoA to ethanol via acetaldehyde is a reversible process (33, 35, 41, 152). During the reaction two molecules of NADH are reoxidized; one is oxidized during the acetyl CoA to acetaldehyde step and the second during the reduction of acetaldehyde to ethanol (Fig. 1). For a long time it was considered that these two reduction steps were catalyzed by distinct enzymes, acetaldehyde dehydrogenase (ACDH) and ADH; however, both genetic and biochemical analyses have clearly demonstrated that a homopolymeric ADH (EC 1.1.1.1) enzyme catalyzes both reactions (66, 99). Recently, the enzyme could be shown to quench specifically the free radical of pyruvate formate-lyase (98, 99; for details, see chapter 15). All three functions of the enzyme are associated with redox processes and are strongly dependent on ferrous iron and pyridine nucleotides (98, 99). Only the PFL-deactivating activity is irreversible, but it clearly requires NAD+ and CoA (98, 99).
Genetics of ADH.
Several screening and selection procedures have been successfully employed to isolate mutants with altered ADH levels. These procedures included looking for ADH-overproducing strains that permitted E. coli to grow on ethanol as sole carbon source (33, 35); mutagenizing ADH-overproducing mutants and screening for loss of ability to grow on ethanol (33, 211); employing suicide substrates, such as chloroethanol, chloroacetaldehyde (39), or allyl alcohol (115), and selecting resistant mutants; even screening for the loss of capacity to reduce the redox dye benzyl viologen (BV), which normally is used to isolate hydrogenase-defective or FHL-defective mutants, proved successful in isolating ADH– mutants (30). All of the mutants lacking ADH enzyme activity were also defective in ACDH activity. Moreover, the mutants were characterized by being unable to ferment hexoses or sugar alcohols. They could only grow anaerobically either by fermenting the more oxidized glucuronate or by respiring with fumarate or nitrate as electron acceptor (30, 33, 35, 72, 99).
The mutants that overproduced ADH also concomitantly overproduced ACDH, thus strengthening the genetic link between the two enzyme activities (33). Mapping studies localized all of the mutations to the 27.3-min region of the chromosome. Finally, cloning of the adhE gene, determination of the DNA sequence, and construction of a specific deletion mutant provided unequivocal proof that both ADH activity and ACDH activity are encoded by a single gene (66, 99). The adhE gene encodes a polypeptide comprising 891 amino acids, with a portion of the polypeptide from amino acids 450 to 850 exhibiting strong similarity to ADH III of Zymomonas mobilis (37) and ADH IV of Saccharomyces cerevisiae (209).
Expression of the adh gene is induced by anaerobiosis. However, this induction is independent of the Fnr and ArcA transcription factors (31, 110). There appears to be a direct correlation between the NADH/NAD+ ratio and enzyme synthesis; the higher the ratio, the more ADH is synthesized (110). Moreover, the ADH protein itself may exert a regulatory function, since gene expression is dramatically enhanced in an adhE mutant (110).
Biochemistry of ADH.
The genetic analysis of ADH has, to a large extent, preceded the biochemical analysis of the enzyme. Apart from an early study (152) that involved the characterization of partially purified aldehyde dehydrogenase activity, the major information on ADH has been secured recently (98, 99). Indeed, it was through the characterization of the PFL-deactivating activity present in anaerobic E. coli cells that the identity of ADH and PFL-deactivase was discovered.
The ADH protein is a homopolymer comprising between 20 and 60 subunits. Each subunit has a molecular weight of 96,000, yielding an M r of several million for the native protein (98, 99). Electron microscopy identified the ADH protein as a left-handed helical rod ranging from 45 to 120 nm in length, most frequently being found with a length of 80 nm. This finding immediately suggested identity between ADH and the "spirosomes" identified in many anaerobic bacteria, including E. coli (123). Moreover, these recent findings are in accord with the observation that aldehyde dehydrogenase activity of E. coli is associated with a large, soluble protein particle (170).
Incubation of purified ADH with NAD+ and Fe2+ ions causes the enzyme structure to extend along its helical axis to a length of 130 nm (98). This implies that a major conformational change occurs within the polymer that is probably associated with its enzymatic functions. As already mentioned, all three activities require Fe2+ ions but the protein also must be able to interact with several substrates; the substrates of the PFL-deactivating reaction are radical-bearing PFL enzyme and NAD+, with a stoichiometric requirement for CoA; ADH activity has either acetaldehyde and NADH or ethanol and NAD+ as substrates; ACDH has acetyl CoA and NADH or acetaldehyde and NAD+ as substrates (98). Where these different enzyme activities are accommodated on this remarkable protein remains to be elucidated; however, Knappe and his colleagues have been able to separate ADH activity physically from the other two enzyme activities by dissecting the enzyme with chymotrypsin (98). Protease treatment cleaved the polypeptide into 86-kDa and 14-kDa polypeptides. The larger peptide retains only ADH activity. Taking into account the similarity between the central portion of ADH with ADHs from other sources, this localizes the ADH activity to a region spanning amino acids 450 to 772 (98). The ACDH and PFL-deactivase activities both have a common CoA requirement, and the N-terminal amino acids 1 to 449, together with the C-terminal portion (amino acids 763 to 891) of the polypeptide, are clearly necessary for these enzyme activities. It is probable that either one or both of these regions also will harbor the polymerization domain of the enzyme (98).
The FHL pathway was first described by Stephenson and Stickland (180, 181). As the name suggests, it catalyzes the disproportionation of formate to carbon dioxide and dihydrogen. Studies in the 1950s identified an absolute requirement of selenium and molybdenum for the synthesis of active formate dehydrogenase (FDH) and hydrogen gas production by E. coli (141), and it was established that an FDH, a hydrogenase, and two electron carriers constituted the FHL pathway (62, 140). The FDH isoenzyme associated with the FHL pathway is termed FDHH, and the hydrogenase 3 isoenzyme is called Hyd-3; both of these enzymes are biochemically and genetically distinct from the other FDH and hydrogenase isoenzymes present in E. coli and S. typhimurium (reviewed in reference 158). According to current evidence, the FHL pathway constitutes a multiprotein complex located on the inner aspect of the cytoplasmic membrane; henceforth it will be referred to as the FHL complex (Fig. 2; 156).
FDHH.
FHL complex activity can be determined as the formate-dependent production of dihydrogen (164), or the activity of the FDHH isoenzyme (EC 1.2.1.–) component can be determined in isolation by measuring the formate-dependent reduction of the one-electron, low-redox-potential dye BV (8, 38, 140). The physiological electron acceptor of FDHH has yet to be determined biochemically, although genetic evidence indicates that it is a protein encoded by the hyc operon (see below).
The FDHH isoenzyme is associated with an 80-kDa selenopolypeptide encoded by the fdhF gene (8, 38, 138, 217). Selenium, in the form of selenocysteine, is located at amino acid position 140 in the FDHH polypeptide chain (179, 216). The FDHH polypeptide has been purified and shown to contain 3.3 g-atoms of iron and 1 g-atom of molybdenum per mol of enzyme (8). These results suggest that the enzyme contains a single 4Fe-4S iron-sulfur cluster. Heider and Bck (78) have recently proposed that a conserved cysteine motif common to Mo-cofactor-dependent FDHs may be involved in liganding the cluster. Molybdenum has been reported to be associated with the enzyme in the form of a molybdopterin guanine dinucleotide cofactor (8). The stoichiometry of FDHH in the FHL complex is not known, mainly because the complex is extremely unstable and so has remained refractory to biochemical analysis. However, it appears that the FDHH component is loosely associated with the complex, since enzyme activity can be readily determined in the soluble fraction after subcellular fractionation of crude extracts (38, 63, 153, 156, 164).
Detailed kinetic studies employing deuteroformate and proteoformate clearly demonstrated that the formate oxidation step is not rate-limiting, but rather the subsequent one-electron transfer steps to BV in the in vitro analyses are rate-determining (7). The direct involvement of the selenol group in formate oxidation was shown by comparing the enzymatic conversion of the selenocysteinyl enzyme with a cysteinyl-substituted derivative. The sulfur enzyme also proved to be more than two orders of magnitude less active than its selenium-containing counterpart at physiological pH, thus emphasizing the advantage of the reactivity of the selenol over the thiol group in redox processes (6).
Hyd-3.
Hyd-3 (EC 1.12.1.–) has proved even more recalcitrant to biochemical analysis than FDHH. The existence of a third hydrogenase isoenzyme was first established by carrying out immunoprecipitation studies with antibodies raised against the Hyd-1 and Hyd-2 isoenzymes (164). The nonimmunoprecipitable hydrogenase enzyme activity could be correlated with FHL synthesis; however, activity was extremely labile and the enzyme could not be purified initially. The identification of the gene (hycE) encoding the large subunit of Hyd-3 facilitated subsequent characterization of the enzyme (21, 156). The HycE polypeptide has been purified and shown to contain up to 1 mol of nickel per mol of enzyme (147). Like FDHH, the HycE polypeptide also appears to form a loose association with the other components of the FHL complex (156). It is likely that both of these polypeptides have an associated small subunit that functions in electron transfer within the complex (Table 3 and see below).
Table 3Function of the fdhF and hyc operon gene products |
Genetics of FHL Complex Formation.
Molecular genetics proved to be the key to unraveling the complexity of the FHL complex. Two mutants were isolated by Pecher and colleagues that were defective in FHL activity (139). One proved to carry a lesion in the fdhF gene encoding the selenopolypeptide of FDHH, while the other had an insertion element located within the second gene of a multicistronic operon (hyc) encoding, among other polypeptides, the other structural components of the FHL complex (21, 138, 139, 215, 217). The fdhF gene is located at 92.6 min, and the hyc operon is located at 58.8 min on the E. coli chromosome. Further DNA sequencing studies around the hyc operon identified a large locus (Fig. 3) required for hydrogenase isoenzyme biosynthesis (21, 117, 168). The hyp operon encodes proteins involved in nickel processing and is required for the synthesis of functional hydrogenase isoenzymes (see below).
A detailed in-frame deletion analysis together with database searches (21, 156) has provided considerable information regarding the putative functions of the hyc operon gene products in the FHL complex (Table 3). The HycA polypeptide is not a structural component of the FHL complex, but rather participates in the transcriptional regulation of complex formation. The exact function of the HycH protein in FHL complex formation is still not clear (Fig. 3).
Transcription of the FHL complex genes occurs only during fermentative growth conditions and is absolutely dependent on formate and the alternative sigma factor NtrA (16, 17, 116, 117, 148). Hence, the expression of the fdhF gene and the hyc and hyp operons is precisely coordinated. The isolation of trans-acting regulatory mutants identified the fhlA gene (Fig. 3) as that encoding the transcriptional regulator which coordinates the expression of these genes in the presence of a critical threshold level of formate (167, 168). The FhlA protein has homology with regulators of two-component sensor-regulator pairs (168). It has been shown to bind specifically to a cis regulatory sequence located approximately 100 bp upstream of the fdhF gene (169), previously characterized by deletion analysis to be essential for the formate-dependent expression of an fdhF-lacZ fusion (15). FhlA binds to two further cis regulatory sequences; one sequence, termed IR1, is located between the hycA and hypA genes (see Fig. 3) of the divergently oriented hyc and hyp operons, while the second binding site (IR2) is located between the hycA and hycB genes (169). Studies employing an in vitro coupled transcription-translation system have demonstrated that IR1 is necessary for activation of hyc operon transcription and IR2 is required to activate transcription of the hyp operon (83). It is still unclear whether formate interacts directly with the FhlA protein to effect transcriptional activation or whether a formate "sensor" protein exists which then relays a signal (for example via a phosphorylation cascade) to FhlA.
The integration host factor has also been shown to be required to optimize the expression from this complex regulatory region. It has been proposed that one function may be to organize a supramolecular transcription complex (83). This factor is not involved in the transcriptional regulation of the fdhF gene.
Several of the hyp operon gene products are also required during respiratory growth conditions (see below). An Inv-dependent promoter is located within the hypA gene to ensure that sufficient levels of these proteins are present for the maturation of catalytically active hydrogenase isoenzymes, even in the absence of formate (117).
The fhlA gene is transcribed at a low level from its own promoter, and this level is enhanced anaerobically through the activity of the Inv-dependent promoter within the hypA gene (see Fig. 3) (148). Activation of the FhlA-dependent promoter in front of hypA further increases fhlA gene transcription. This scenario presents a novel positive-feedback mechanism for transcriptional control of a regulon. The HycA protein appears to antagonize the action of FhlA, thus preventing persistent activation of the formate regulon (S. Hopper and A. Bck, unpublished data). Exactly how HycA achieves this is still unclear.
Physiology of Dihydrogen Production.
The intracellular formate concentration is the crucial determinant of FHL complex biosynthesis (148). When E. coli grows at neutral pH, formate is exported from the cell and the FHL complex is synthesized at very low levels (14, 148, 189). As the extracellular concentration of the fermentation products acetate, lactate, succinate, and formate increases, there is a concomitant decrease in the pH of the medium. As a means of offsetting this pH drop, formate is reimported into the cell (148, 189), attains a critical threshold concentration,and activates the transcriptional regulator FhlA, which consequently induces FHL complex biosynthesis (148, 168; see chapter 95). The dihydrogen produced can then either diffuse away from the cell or be reoxidized by Hyd-1 or Hyd-2 and used as a source of energy. The FHL complex therefore is a means of maintaining pH homeostasis.
The Enzymes.
It is not possible to distinguish Hyd-1, Hyd-2, or Hyd-3 on the basis of enzyme activity determination because the assay for all three measures the dihydrogen-dependent reduction of BV. The isoenzymes are, however, distinguishable immunologically (9, 164). Both Hyd-1 and Hyd-2 (EC 1.12.1.–) have been purified (2, 10, 58, 165), and their properties are summarized in Table 4. Hyd-2 can be purified as a soluble, active, tryptic fragment that differs from the native membrane-bound enzyme only through the loss of a 5-kDa fragment from the small subunit (10). The data indicate that the enzyme is anchored in the membrane but the large proportion of the enzyme is exposed at the periplasmic surface of the membrane. In contrast, Hyd-1 cannot be released from the membrane fraction by proteolysis, but it can be readily solubilized by detergents such as Triton X-100 (9, 164, 165).
Table 4Comparison of the properties of the hydrogenase isoenzymes |
Neither purified Hyd-1 nor Hyd-2 reacts with quinones, although the quinone pool might be expected to receive the electrons derived from dihydrogen oxidation catalyzed by these enzymes (see below). This suggests that a further subunit involved in electron transfer to the quinone pool may have been lost during purification (44, 158). Both enzymes have a very low apparent Km for dihydrogen, and this is in accord with the enzymes’ function in H2 oxidation (10, 165). Hyd-2 has a greater capacity for dihydrogen oxidation than Hyd-1 (10).
Genetics of Hyd-1 and Hyd-2.
Mutants specifically defective in Hyd-2 biosynthesis have been isolated (102, 106, 182). The mutations are likely to be located within the structural genes of the hyb operon, which encodes Hyd-2 (142). The operon comprises seven genes located at 65 min on the E. coli chromosome (Table 5). Three of the genes, hybA, hybB, and hybC, encode structural components of the enzyme; the designation of hybB as a subunit is based on sequence homologies with the third subunit of the hydrogenase of Wolinella succinogenes (44). The other four hyb genes, hybD, hybE, hybF, and hybG, are essential for synthesis of a fully active Hyd-2 isoenzyme, but their exact functions are still obscure (142).
Table 5Function of the gene products of the hya and hyb operons |
The structural genes of Hyd-1 are encoded by the hya operon located at 22.1 min on the E. coli chromosome (127, 128, 142). The hya operon has six genes, with hyaA, hyaB, and hyaC encoding the subunits of the enzyme (Table 5). The hyaC gene product exhibits homology with HybB and probably has a similar function. Like the accessory polypeptides encoded by the hyb operon, the hyaD, hyaE, and hyaF gene products are essential for synthesis of functional Hyd-1 (128, 142). The large and small subunits of both Hyd-1 and Hyd-2 share extensive similarities with the respective hydrogenase polypeptides from other organisms. The various implications these homologies may have with regard to the structure and function of hydrogenases in general have been reviewed in detail (59, 142, 204, 212).
Physiology of Hyd-1 and Hyd-2.
Although Hyd-1 and Hyd-2 are present at substantial levels when E. coli or S. typhimurium cells ferment hexoses (87, 164, 166), it is still questionable whether they can be classified as true enzymes of fermentation. It is clear that Hyd-2 is the principal H2-oxidizing activity when E. coli cells grow on dihydrogen and fumarate (164, 166), and the enzyme is probably proton translocating (94). Hyd-1 spans the cytoplasmic membrane and is probably also an energy-conserving enzyme (67). The likely topologies of the enzymes in the cytoplasmic membrane are also in agreement with their H2-oxidizing capacities (Fig. 2). However, under which particular growth conditions Hyd-1 oxidizes dihydrogen is still open to question (158).
It has been proposed that either Hyd-1 or Hyd-2, or both, could serve the function of recycling the dihydrogen evolved by the FHL complex during fermentation (164, 166). Such an H2-recycling mechanism could be useful in facilitating redox balance, for example when particularly reducing substrates such as sugar alcohols are oxidized in the absence of exogenous electron acceptors (34). Evidence has been presented which indicates that when growing on sorbitol, fermenting E. coli cells produce excess ethanol and increased amounts of succinate relative to acetate and formate, which cannot be accounted for by calculating redox balance using standard fermentation pathways (3). One means of accounting for the excess ethanol production would be if the reducing equivalents from formate were recycled and channeled to fumarate via the quinone pool (Fig. 2). This could be achieved by Hyd-1- or Hyd-2-dependent reoxidation of some of the dihydrogen produced by the FHL complex. Indeed, Alam and Clark showed that in a hypB mutant, which is incapable of synthesizing any of the Hyd isoenzymes, the amount of ethanol and succinate produced was significantly decreased (3). Substantiation of these results using specific deletion mutants may strengthen the proposal that Hyd-1 and Hyd-2 can function in fermentation.
The formation of many of the enzymes involved in fermentation reactions depends on the availability, the uptake, and the incorporation of a number of metals and metal-containing cofactors. The major ones involved are Fe, Mo, Ni, Co, and Se. The acquisition of iron, molybdenum, and cobalt and the biosynthesis of the molybdenum-containing cofactor and of coenzyme B12 are described elsewhere in this volume (chapters 47 and 71). The metabolism of selenium and of nickel, however, is intimately, and almost exclusively, associated with fermentation reactions. Therefore, an account of it will be presented here.
The requirement of selenium for the degradation of formate was recognized as early as 1954 (141), but it took more than 30 years before the major features of its metabolism were resolved (for reviews see references 20, 28, 78, 177, and 178). Selenium is incorporated into three FDH isoenzymes in the form of a single selenocysteine residue. The identity of two of these isoenzymes was disclosed by labelling with the 75Se radioisotope (38). Fermentative growth of E. coli on glucose as carbon source resulted in the specific labeling of an 80-kDa selenopolypeptide which correlated with the formation of FDHH enzyme activity (38, 138). Growth under nitrate respiratory conditions labeled a 110-kDa selenopolypeptide which is a subunit of FDH N (FDHN). The enzyme delivers the electrons withdrawn from formate to nitrate reductase (38, 48). A third FDH isoenzyme (FDHO) is present in cells grown in the presence of oxygen or anaerobically in the presence of nitrate (161). It possesses a 110-kDa selenopolypeptide and appears to feed the electrons derived from formate into either the aerobic or the nitrate respiratory chain. Its actual physiological role, however, has yet to be resolved. Apart from being a constituent of the selenopolypeptides of the FDHs, selenium is also incorporated into modified nucleosides of some tRNAs (28, 177).
Mutants (termed fdh) had been isolated previously that were pleiotropically defective in the activity of the two FDHs known to exist at that time, FDHH and FDHN (Table 6). A reisolation and detailed analysis of several classes of mutants disclosed that they carried a lesion in one of the steps of biosynthesis or incorporation of selenocysteine (108). The selC gene codes for a unique tRNA (tRNASec) which is charged by seryl tRNA-synthetase (Fig. 4) (109). Seryl-tRNASec is then bound to selenocysteine synthase (the selA gene product) via a Schiff base between the amino group of the seryl residue and the carbonyl group of the pyridoxal 5'-phosphate of the enzyme (52, 54). The 2,3- elimination of a water molecule yields enzyme-bound dehydroalanyl-tRNASec. Addition of reduced and activated selenium to the double bond results in the liberation of selenocysteyl-tRNASec from the enzyme. The activated selenium compound is a product of the reaction catalyzed by the selD gene product (selenophosphate synthetase) (107). This enzyme cleaves both anhydride bonds of ATP, thereby transferring the γ-phosphate to selenide, which results in selenophosphate synthesis and the release of AMP and of the β-phosphate of ATP (46, 203).
Table 6Gene products involved in selenocysteine biosynthesis and incorporation |
Selenocysteine synthase (EC 4.2.1.–) is a homodecamer of 50-kDa constituent subunits (54). High-resolution electron microscopy revealed that the decamer is able to bind five seryl-tRNASec molecules (47). A stoichiometry of two enzyme subunits to one tRNA molecule could also be demonstrated by biochemical analyses (52).
Selenophosphate synthetase (the selD gene product; EC 6.5.1.–) is a 37-kDa monomer (107). It possesses a conspicuous –Cys-17–Gly–Cys-19– motif close to the N terminus of the protein, and it was shown that the cysteinyl residue at position 17 is essential for catalytic activity (100).
Selenocysteyl-tRNASec requires a specialized translation factor, SELB, for the decoding process at the ribosome. SELB is homologous in its function to EF-Tu and therefore binds guanine nucleotides and the charged tRNASec (Fig. 4) (53). However, in contrast to EF-Tu it can discriminate between aminoacyl residues. Therefore, seryl-tRNASec is not a substrate for the protein. Neither seryl-tRNASec nor selenocysteyl-tRNASec is recognized by EF-Tu to a physiologically significant extent (55). SELB also has the capacity to bind to a specific recognition motif which is present in mRNAs coding for prokaryotic selenoproteins and which is located immediately downstream of the UGA directing selenocysteine insertion (12, 76, 77, 218). The decoding process is reviewed in this volume (chapter 60).
The flux into and the fidelity of selenocysteine biosynthesis and incorporation are regulated at several levels. First, seryl-tRNA synthetase charges tRNASec with only about 1% of the catalytic efficiency compared to tRNASer, which reflects the very minor amount of serine carbon required for selenocysteine synthesis (11, 112). Second, tRNASec possesses structural features which discriminate it from tRNASer in its reaction with selenocysteine synthase (52, 109) translation factor SELB and in its incompatibility of reacting with EF-Tu (11, 13, 55, 112). Finally, SELB binds tRNASec only when it is charged with selenocysteine; seryl-tRNASec, the precursor, is not recognized (53).
The exact biochemical function of the selenocysteine residue in catalysis has not been resolved yet. A cysteine-containing variant has been constructed genetically and purified (6). It was found to be active catalytically but with a K cat 2 orders of magnitude less than the Se form (6, 8). Also, naturally occurring S forms homologous to the E. coli FDHs have been characterized (22, 172). Since they are abundant proteins in these organisms, the selective advantage of having a selenol in the active site might be a means of lowering the amount of enzyme required to fulfil a particular catalytic task (28, 177).
The sel genes are expressed constitutively in E. coli, probably reflecting the need for synthesis of the selenopolypeptides under aerobic (FDHO), nitrate respiratory (FDHN and FDHO), and fermentative (FDHH) conditions (161).
The three hydrogenase isoenzymes from E. coli belong to the class of [NiFe] hydrogenases (9, 164). According to current models, the nickel appears to be liganded to the large subunit of these hydrogenases, with the possible involvement of two highly conserved motifs, RXCXXC close to the N terminus and DPCXXCXXH at the C terminus (142, 144).
Ni2+ ions are taken up from the medium by at least two active transport systems, (i) a high-capacity magnesium uptake system that is rather unspecific and can recognize and transport nickel and (ii) a high-specificity but low-capacity nickel transport system (75). The former is encoded by the corA gene (136), and the latter is encoded by the five genes of the nik operon (130). According to the derived amino acid sequences, the products of the nik operon belong to the class of periplasmic-binding-protein- dependent uptake systems, the so-called ABC transporters (80). Expression of the nik operon genes is repressed by high nickel in the medium and is under the positive control of the fnr gene product (213, 214). Such fnr mutants, therefore, are devoid of hydrogenase activity in the presence of high magnesium ion concentrations, i.e., when Ni uptake by the CorA-dependent system is out-competed by Mg2+ ions.
Genetic analysis of hydrogenase formation in E. coli revealed the existence of a number of genes (hyp) whose products have a function in the formation of hydrogenase activity rather than in its synthesis (117). They are encoded in the hyp operon (hypA–E) (Fig. 3) and by a gene located downstream of the hyc operon (hypF) (96). All of these gene products are required for synthesis of active Hyd-3, while lesions in some of the genes also affect the biosynthesis of the Hyd-1 and Hyd-2 isoenzymes (86). Thus, mutants carrying lesions in hypB, hypD, hypE, and hypF lack all three hydrogenase isoenzymes, indicating that all of these gene products are required for the maturation of catalytically active hydrogenase isoenzymes in E. coli (Fig. 3). hypA mutants, on the other hand, have unimpaired levels of Hyd-1 but greatly increased Hyd-2 activity. hypC mutations do not affect Hyd-2 levels but result in decreased Hyd-1 activity (86). The differential effects of hypA and hypC mutations on the development of activity of the three hydrogenase isoenzymes may be due to the existence of homologous genes encoded by other operons. For example, the hyb operon encoding Hyd-2 contains a gene (hypG) whose product exhibits strong amino acid sequence similarity to the hypC gene product (142). This could be the basis of why hypC mutations are specific for Hyd-3 only but have a partial effect on Hyd-1 synthesis (Fig. 3).
The catalytic functions of the hyp gene products in hydrogenase isoenzyme maturation are largely unknown. Possible functions range from nickel insertion, protein folding, proteolytic processing, and membrane integration, to reductive activation (117, 142).
A particularly intriguing protein is the hypB gene product. Mutations in hypB can be phenotypically suppressed by high nickel concentrations in the medium (86, 117). The hypB mutants, therefore, have a phenotype similar to that exhibited by fnr mutants on the expression of the nik operon. Despite this phenotypic similarity, HypB must serve a distinct function, since it is localized in the cytoplasm (120) and hypB mutants are able to accumulate nickel to an extent that active urease from Proteus vulgaris can be synthesized in E. coli (cited in reference 86).
As predicted by an analysis of the amino acid sequence, HypB binds guanine nucleotides (120). A model has been proposed which suggests that HypB is a Ni-binding protein which donates the metal to the hydrogenase apoprotein. GTP binding and hydrolysis may be involved in releasing HypB from the apoprotein once the metal has been released (120).
A common property of the hyp mutants is that the large subunit of the hydrogenases is present in an unprocessed form which does not contain nickel (86, 117). Processing involves the C-terminal cleavage and the release of a 32-amino-acid polypeptide in the case of the large subunit (HycE) of Hyd-3 (147). The site of cleavage is at the C-terminal side of the Arg-537 residue of this protein. Large subunits of all other hydrogenases analyzed thus far have a His residue in this position (65, 142). It remains to be determined whether this deviation from the consensus in Hyd-3 is correlated with its function as a gas-evolving enzyme.
The proteolytic processing could be reconstituted in an in vitro reaction using purified components (147). Proteolysis occurs only when the precursor has been purified from a strain possessing a deletion in the hycH gene. In contrast to the situation in hyp mutants, this precursor contains nickel (147), indicating that for C-terminal processing to occur it is conditional that nickel has to be coordinated. The exact function of the hycH gene product in FHL complex assembly is still open; it is not identical with the protease required for the processing step (147).
In much of the early literature, reference is made to "Aerobacter" isolates. This was a general term that was used until the end of the 1950s to classify many Aerobacter and Klebsiella species because at that time it was not possible to distinguish categorically one from the other (23). In 1960 the genus Enterobacter was proposed (84), and by employing various biochemical differentiation criteria, and more recently with the aid of DNA/RNA analyses, it has become possible to classify these organisms with relative ease.
Some of the more commonly found synonyms are listed in Table 7. The discovery of amino acid decarboxylases aided greatly in the differentiation procedure (24, 25, 68, 69). Thus, Enterobacter aerogenes and Klebsiella pneumoniae are readily distinguished on the basis of their ornithine decarboxylase phenotype; the former organism is positive and the latter is negative. Aerobacter indologenes (146) has been classified as Klebsiella oxytoca due to its indole production (Table 7). Both K. pneumoniae and Enterobacter aerogenes are indole negative.
Table 7Nomenclature of the family Enterobacteriaceae |
The Pathway and the Enzymes.
In 1906, Harden and Walpole (74) discovered that the product pattern of fermenting Aerobacter aerogenes differs from that of E. coli due to the presence of several neutral compounds which were later identified as 2,3-butanediol and acetoin. The production of acetoin (in its oxidized form as diacetyl) is the basis of one of the most frequently used identification tests in bacteriology, the Voges-Proskauer reaction (45). The capacity to produce 2,3-butanediol is widely distributed among microorganisms. Typical producing species are found in the genera Klebsiella, Enterobacter, Serratia, Bacillus, Lactobacillus, and Aeromonas. In none of these organisms is 2,3-butanediol the sole fermentation product. These organisms can be superficially classified into diol-H2 producers (K. pneumoniae; Aeromonas hydrophila; Bacillus polymyxa), diol-formate producers (Serratia marcescens), and diol-glycerol producers (Bacillus subtilis), based on the end product whose yield is second to butanediol (119). It should be stressed that the pathway described for enterobacteria is not valid for all organisms that exhibit a positive reaction in the Voges-Proskauer test.
Table 8 compares the fermentation balance of K. oxytoca (Aerobacter indologenes) (146) with that of E. coli (18). E. coli produces only traces of 2,3-butanediol and acetoin whereas the diol is one of the major products of K. oxytoca.
Table 8Comparison of the fermentation products of E.coli and K.oxytoca (Aerobacter indologenes) |
Three enzymes are involved in the conversion of 2 mol of pyruvic acid into 1 mol of 2,3-butanediol (Fig. 5): α-acetolactate synthase (ALS), α-acetolactate decarboxylase (ALDC), and acetoin reductase (AR).
ALS.
ALS (EC 4.1.3.18) is a thiamine PPi-containing enzyme. One molecule of pyruvate forms an adduct with the thiamine PPi and is then decarboxylated, leaving a hydroxyethyl–thiamine PPi–enzyme intermediate. The resonance-stabilized carbanion can attack the keto group of a second molecule of pyruvate, delivering α-acetolactate (Fig. 5).
ALS has been purified by Strmer and coworkers from both Aerobacter aerogenes (185) and Serratia marcescens (122). The enzyme from these organisms exhibits a narrow pH optimum of activity around 6.0 and has been denoted as the "pH 6.0" acetolactate-forming enzyme, or catabolic ALS, to differentiate it from the α-acetohydroxy-acid synthases which are most active at alkaline pH (7.8 to 8.0) and serve an anabolic function in isoleucine and valine biosynthesis (see Fig. 5). ALS is a homodimer comprising 58-kDa constituent subunits (85).
Stationary-phase cells of B. subtilis have been shown to synthesize a catabolic ALS which, in this particular case, displays a pH optimum of activity around 7.0 (82).
ALDC.
ALDC (EC 4.1.1.5) is the most thoroughly investigated enzyme of the pathway. It has been purified from Aerobacter aerogenes (114), Lactobacillus casei (143), Brevibacterium acetylicum (135), and Bacillus brevis (191). The pH optimum of the Aerobacter enzyme is also in the acidic range, between 6.2 and 6.4. ALDC, as ALS, is a functional dimer (114). The occurrence of the enzyme appears to be restricted to prokaryotes (64).
AR.
AR (EC 1.1.1.5) catalyzes the reduction of acetoin to 2,3-butanediol with NADH as reductant. The enzyme has been purified from Aerobacter aerogenes and shown to be a homotetramer made up of 25-kDa subunits (186). The acetoin reduction reaction is reversible, so under conditions of a high NAD+/NADH ratio AR can act as a butanediol dehydrogenase. Interestingly, AR can also reduce diacetyl to acetoin but in an irreversible fashion. The pH optimum of activity depends on the substrate; it lies between pH 4 and pH 7 with diacetyl or acetoin as substrate and is at 9.5 with 2,3-butanediol as substrate (186). There are three stereoisomeric forms of 2,3-butanediol which are all produced to different extents by the different microorganisms: the meso, d-(–), and l-(+) forms. K. pneumoniae has been shown to produce between 5 and 14% l-(+) and the remainder meso-2,3-butanediol (for a review see reference 82). There has been much debate about the formation of these stereoisomers, and different models have been proposed (82). The one valid for the Klebsiella situation may involve the activity of an l-(+)-specific AR responsible for the formation of the minor amount of l-(+)-2,3-butanediol. Since acetoin is produced by ALDC in the d-(–) form, the existence of a racemase must be postulated to account for the interconversion of d-(–)- and l-(+)-acetoin (82). Evidence for an l-(+)-specific AR in Klebsiella comes from the finding that this organism is unable to oxidize d-(–)-butanediol (81).
Physiology of 2,3-Butanediol Formation.
2,3-Butanediol formation by Enterobacter and Klebsiella spp. requires anaerobiosis, an acidic pH, and the presence of acetate in the culture medium (91, 184, 187). Acetate appears to play the key role in the regulation of the activity and the formation of the pathway, and it is possible, albeit not proven, that anaerobiosis and low pH are mechanistically connected with the effect of acetate. Thus, the entry of acetate may be favored by low pH, and anaerobiosis might simply increase the rate of acetate formation from glucose.
Figure 6 illustrates the current concept of how acetate may stimulate 2,3-butanediol formation in Enterobacter and Klebsiella spp. and is based on studies by Strmer and coworkers (91, 184, 187). Acetate stimulates the activity of ALS and inhibits oxidation of 2,3-butanediol to acetoin at the enzyme activity level. Moreover, formation of the pathway is strongly dependent on the presence of acetate in the medium. A systematic study of the expression and activity of the 2,3-butanediol pathway by Enterobacter aerogenes during growth in batch culture on glucose revealed that the pH of the medium initially dropped to 5.8 as a consequence of acetate accumulation. The low pH resulted in growth cessation and in acetoin and 2,3-butanediol formation. When glucose became exhausted the pH increased to 6.5, and this was accompanied by some reoxidation of 2,3-butanediol to acetoin (91). This time course may reflect the following physiological scenario. Acetate is produced (and ATP is generated by PTA-ACK) until the pH becomes limiting. Acetate (or a metabolic product thereof) then induces the expression of the 2,3-butanediol pathway and more than 50% of the pyruvate is channeled into the formation of the neutral end product with concomitant NAD+ regeneration. When the glucose becomes exhausted, 2,3-butanediol can serve as a reservoir of reducing equivalents (NADH) (91, 119). The genes coding for the three enzymes of the 2,3-butanediol pathway in Klebsiella terrigena and Enterobacter aerogenes have been characterized recently (19). They form an operon (budABC) in which budA codes for the ALDC enzymes and budB and budC encode ALS and AR, respectively. No regulatory gene has been identified either in Klebsiella or in Enterobacter.
Such a regulatory gene, alsR, however, has been recently identified in B. subtilis (145). Inactivation of alsR leads to the loss of expression of the alsSD operon in which alsS codes for ALS and alsD codes for ALDC of the 2,3-butanediol production pathway. As in enterobacteria, expression appears to be induced by acetate (82).
The Pathway.
Glycerol is a constituent component of lipids and fats and, therefore, is an abundant carbon and energy source for microorganisms in nature. In the presence of exogenous electron acceptors, glycerol is utilized exclusively via a pathway comprising an ATP-dependent kinase and, depending on the type of exogenous electron acceptor, one of two flavin-linked sn-glycerol-3-phosphate dehydrogenases. Dihydroxyacetone phosphate is the product of this pathway and enters glycolysis via the triose-phosphate isomerase reaction (chapter 20). This pathway is present in all members of the Enterobacteriaceae examined to date.
Fermentative growth on glycerol as a carbon source, however, is much more restricted among microorganisms. This may reflect the fact that glycerol (per average carbon atom) is more reduced than carbohydrates. Only a few species of the enterobacteria are able to degrade it in the absence of an exogenous hydrogen acceptor, and these are principally members of the genera Klebsiella and Citrobacter. This higher reduction state of glycerol is overcome in these organisms by the cooperative action of two pathways (Fig. 7). An oxidative pathway is constituted by a NAD+-dependent glycerol dehydrogenase, which yields dihydroxyacetone from glycerol, and an ATP-dependent dihydroxyacetone kinase (113, 118, 154). The resulting dihydroxyacetone phosphate is then channeled into intermediary metabolism.
The reductive branch consists of a coenzyme B12-dependent glycerol dehydratase, forming β-hydroxypropionaldehyde from a second molecule of glycerol, and a 1,3-propanediol dehydrogenase which reoxidizes the NADH generated in the oxidative branch (1). As a consequence, glycerol is either oxidized or reduced, yielding nearly equimolar amounts of dihydroxyacetone phosphate and 1,3-propanediol, respectively; the latter is excreted into the medium (Fig. 7). A significant amount of acrolein is also formed in the course of glycerol fermentation by Klebsiella spp. This could be ascribed to the spontaneous dehydration of β-hydroxypropionaldehyde (1).
Glycerol Dehydrogenase.
Glycerol dehydrogenase (glycerol:NAD+ oxidoreductase, EC 1.1.1.6) catalyzes the reaction: glycerol + NAD+ → dihydroxyacetone + NADH + H+. One of the prominent properties of the enzyme is its activation by monovalent cations, which was first described by Lin et al. for the enzyme of an encapsulated strain of Klebsiella pneumoniae 1033 (113). This feature of the enzyme has been intensively investigated by McGregor and colleagues with a partially purified preparation from the same organism (126). In the absence of cations, activity of glycerol dehydrogenase is reduced to 25 to 30% of that of a fully saturated enzyme. The presence of monovalent cations greatly affects the enzyme’s affinity for the substrates glycerol and dihydroxyacetone, which are increased 22-fold and 25-fold, respectively, at pH 7.5, and the Vmax of the enzyme, which is increased 3-fold. In contrast, neither the affinity for NAD+ nor the ordered bi-bi kinetic mechanism is affected by cations. The order of the efficacy of the cations in the activation was shown to be NH4 + > TI+ > K+ > Rb+ and was independent of pH (126).
Glycerol dehydrogenase has been purified to apparent homogeneity and shown by gel electrophoresis to have a molecular weight of 79,000 (151). Two different aggregation states of the enzyme appear to exist, depending on the pH: an active homodimer composed of M r-40,000 subunits at pH 8.6 and a homotetramer of 160,000 molecular weight at pH 7.0 (151).
Dihydroxyacetone Kinase.
Dihydroxyacetone kinase (EC 2.2.1.3) is a cytoplasmic enzyme that catalyzes the reaction dihydroxyacetone + ATP → dihydroxyacetone phosphate + ADP. The enzyme has been purified from a mutant of Klebsiella pneumoniae 1033 which expresses the dha regulon (see below) constitutively (92). The enzyme is a homodimer of 53,000-molecular-weight subunits and exhibits a high specificity for dihydroxyacetone. Glycerol is not a substrate for the enzyme, nor does it inhibit the enzymic reaction, even at concentrations as high as 0.1 M. Glycerol kinase, on the other hand, accepts both glycerol and diydroxyacetone as substrates. Dihydroxyacetone kinase is not allosterically inhibited by fructose 1,6-bisphosphate, but glycerol kinase is inhibited by this metabolite. Furthermore, immunological cross-reaction could not be detected between the two kinases, supporting the contention that they are distinct enzymes (92).
Glycerol Dehydratase.
In the reductive branch of glycerol fermentation, glycerol is first oxidized by a coenzyme B12-dependent diol dehydratase with the resultant elimination of a water molecule (Fig. 7). Klebsiella species have been reported to possess two different types of coenzyme B12-dependent diol dehydratases: diol dehydratase (d,l-propane-1,2-diol hydrolyase; EC 4.2.1.28) and glycerol dehydratase (glycerol hydrolyase; EC 4.2.1.30). They may occur either singly or together in the same species of Enterobacteriaceae (49, 50, 200). The activity of both dehydratases can be measured in the same extract, since they differ 67-fold in their affinity for coenzyme B12 (49). Both enzymes catalyze the dehydration of ethane 1,2-diol, propane 1,2-diol, and glycerol to acetaldehyde, propionaldehyde, and 3-hydroxypropionaldehyde, respectively. Diol dehydratase formation is induced in K. pneumoniae ATCC 25955 and Citrobacter freundii NCIB 3735 upon anaerobic growth in the presence of propane 1,2-diol, whereas anaerobic growth at the expense of glycerol induces the formation of both enzymes in strain ATCC 25955 (49). Interestingly, K. pneumoniae 8724 forms only diol dehydratase and strain 418 forms only glycerol dehydratase upon growth on glycerol, indicating that both enzymes can function physiologically in the formation of the electron sink 3-hydroxypropionaldehyde (49). Although diol dehydratase may thus substitute for the function of glycerol dehydratase in glycerol fermentation, its actual physiological role has not yet been resolved.
Diol dehydratase and glycerol dehydratase, first purified by Stroinsky et al. (188), have similar molecular weights of about 1.9 105 (49). Both holoenzymes are oxygen sensitive and both are inactivated by their substrate, glycerol, during the course of the enzyme reaction (49, 199, 206).
Propane-1,3-Diol Dehydrogenase.
Propane-1,3-diol dehydrogenase (EC 1.1.1.202) catalyzes the reaction: 3-OH-propionaldehyde + NADH → propane-1,3-diol + NAD+. It has not yet been purified and characterized.
E. coli
and S. typhimurium versus Klebsiella and Citrobacter.
E. coli and S. typhimurium possess the glp system of glycerol degradation but they are devoid of the genes of the dha regulon. They are, therefore, unable to grow anaerobically on glycerol in the absence of an external electron source. Intriguingly, however, these organisms possess a glycerol dehydrogenase that is encoded by the gldA gene at 89.2 min on the E. coli chromosome (201). The enzyme was first purified by Asnis and Brodie (5) from a wild strain of E. coli and subsequently from a mutant producing high, constitutive levels of the enzyme (192). The purified enzyme exists as a homodimer and as a homooctamer of 39-kDa subunits, with both forms displaying enzyme activity (192). The enzyme is activated by monovalent cations, it has a broad substrate specificity, and its formation is induced by hydroxyacetone in the stationary phase of growth (42, 97, 192, 201).
Considerable efforts have been expended on constructing E. coli strains with engineered glycerol degradation pathways. Starting with a glpD glpR double mutant which does not grow on glycerol-based medium aerobically (possibly because of the accumulation of glycerol 3-phosphate), St. Martin and colleagues used mutagenesis and selection for growth on glycerol to isolate a strain which had lost glycerol kinase activity and gained the capacity to grow on high glycerol concentrations (176). Further selection for (aerobic) growth on low glycerol concentrations yielded a mutant with an elevated level of glycerol dehydrogenase. Unfortunately, this strain was still unable to grow on glycerol fermentatively, almost certainly because of the lack of a hydrogen acceptor. Nevertheless, these studies identified an inducible PTS specific for dihydroxyacetone (89).
A further study attempted to generate a strain with a replacement of the glp degradation route for glycerol by a novel pathway. It was hoped that a normally cryptic pathway for glycerol degradation might be induced. This approach, however, resulted in a strain with elevated levels of both glycerol dehydrogenase and the dihydroxyacetone-specific enzyme II of the PTA (90).
Physiology of Glycerol Fermentation.
Growth studies and the isolation and characterization of mutants of K. pneumoniae that were constitutive in the formation of enzymes of the glp regulon, of the glycerol fermentation pathway, and of the diol dehydratase demonstrated that these three systems belong to independent regulatory circuits (50, 51, 149, 150). The rate of formation of the four specific enzymes of the glycerol fermentation pathway responds to the level of dihydroxyacetone, and therefore the gene designation dha was adopted (50). The dhaD gene encodes glycerol dehydrogenase, dhaK encodes the dihydroxyacetone kinase, dhaB encodes the glycerol dehydratase, and dhaT encodes the propane-1,3-diol dehydrogenase. Expression of the dha genes is controlled by the product of the regulatory gene dhaR (50, 51). Furthermore, it is subject to cyclic AMP-mediated catabolite control.
The organization of the dha genes in Klebsiella and Citrobacter spp. has not been resolved to date. Conjugative mobilization of the capacity for fermentative growth of glycerol into E. coli and the demonstration that products of the dhaD, dhaK, and dhaT genes are formed in the transconjugants had indicated that they may be clustered on the Klebsiella chromosome (175). Additionally, induction of their expression by dihydroxyacetone pointed to a cotransfer of the dhaR gene. Active dhaB gene product was not present in the transconjugants. This was interpreted to be the consequence of the inability of E. coli to synthesize coenzyme B12, or else to reflect the fact that the dhaB gene is not genetically linked with the other genes. The absence of glycerol dehydratase in the transconjugants clearly is the basis for the observed lack of formation of propane 1,3-diol (175). It is still unclear how these cells dispose of their reducing equivalents during growth on glycerol in the absence of exogenous electron acceptors.
With the use of a cosmid library from C. freundii, Daniel and Gottschalk recently were able to transform the complete dha regulon into E. coli (40). Expression of active dhaD gene product was shown to be dependent on the presence of a corrinoid in the medium. It was also found that the formation of the enzymes was optimal at low temperature (below 30C); almost no synthesis took place at 37C. The transformants produced propane-1,3-diol when the medium was supplemented with a corrinoid (40).
Shifting a fermenting culture of Klebsiella sp. to aerobic conditions leads to a relatively rapid cessation of the flux through the glycerol pathway. This was shown to be due to the inactivation of glycerol dehydrogenase and also of propane-1,3-diol dehydrogenase (32, 93, 113, 151). Inactivation occurred with a half-life of 45 min in vivo. Inactivation of the enzyme in vitro required the presence of oxygen and a reducing agent. This was thought to result in the formation of hydrogen peroxide and subsequently, in the presence of Fe2+, of a reactive oxygen derivative that attacked the enzyme (93). Indeed, it could be shown that addition of low concentrations of hydrogen peroxide to a fermenting culture led to the inactivation of glycerol dehydrogenase and propane-1,3-diol dehydrogenase (32). Active protein synthesis was required for inactivation by oxygen but not by hydrogen peroxide, suggesting that de novo synthesis of a particular protein is required for the formation of hydrogen peroxide in vivo.
The inactivated (i.e., oxidized) protein had about 10% of the catalytic activity of the native form and was indistinguishable from it in amino acid composition, Km , and molecular weight. It was, however, much more amenable to degradation by proteases, for example (32).
Rapid inactivation of glycerol dehydrogenase and the B12-dependent dehydratase upon a shift to aerobic conditions presumably reduces the concentration of the inducer dihydroxyacetone, guaranteeing a rapid onset of repression. The inactivation, in addition, leads to an immediate block of glycerol degradation via the dha system and promotes a shift to the use of the glp degradation system (51).
The work carried out in the authors’ laboratories has been supported by the Deutsche Forschungsgemeinschaft and the Bundesministerium fr Forschung und Technologie.
References
1. Abeles, R. H., A. M. Brownstein, and C. H. Randles. 1960. β-Hydroxypropionaldehyde, an intermediate in the formation of 1,3-propanediol of Aerobacter aerogenes. Biochim. Biophys. Acta 41:530–531.
2. Adams, M. W. W., and D. O. Hall. 1979. Purification of membrane-bound hydrogenase of Escherichia coli. Biochem. J. 183:11–22.
3. Alam, K. Y., and D. P. Clark. 1989. Anaerobic fermentation balance of Escherichia coli as observed by in vivo nuclear magnetic resonance spectroscopy. J. Bacteriol. 171:6213–6217.
4. Anthony, R. S., and L. B. Spector. 1970. A phosphoenzyme intermediary in acetate kinase action. J. Biol. Chem. 245:6739–6741.
5. Asnis, R., and A. Brodie. 1952. A glycerol dehydrogenase from Escherichia coli. J. Biol. Chem. 203:153–159.
6. Axley, M. J., A. Bck, and T. C. Stadtman. 1991. Catalytic properties of an Escherichia coli formate dehydrogenase mutant in which sulfur replaces selenium. Proc. Natl. Acad. Sci. USA 88:8450–8454.
7. Axley, M. J., and D. A. Grahame. 1991. Kinetics of formate dehydrogenase of Escherichia coli formate-hydrogenlyase. J. Biol. Chem. 266:13731–13736.
8. Axley, M. J., D. A. Grahame, and T. C. Stadtman. 1990. Escherichia coli formate-hydrogen lyase: purification and properties of the selenium-dependent formate dehydrogenase component. J. Biol. Chem. 265:18213–18218.
9. Ballantine, S. P., and D. H. Boxer. 1985. Nickel-containing hydrogenase isoenzymes from anaerobically grown Escherichia coli K-12. J. Bacteriol. 163:454–459.
10. Ballantine, S. P., and D. H. Boxer. 1986. Isolation and characterisation of a soluble active fragment of hydrogenase isoenzyme 2 from the membranes of anaerobically grown Escherichia coli. Eur. J. Biochem. 156:277–284.
11. Baron, C., and A. Bck. 1991. The length of the aminoacyl-acceptor stem of the selenocysteine-specific tRNASec of Escherichia coli is the determinant for binding to elongation factors SELB or Tu. J. Biol. Chem. 266:20375–20379.
12. Baron, C., J. Heider, and A. Bck. 1993. Interaction of translation factor SELB with the formate dehydrogenase H selenopolypeptide mRNA. Proc. Natl. Acad. Sci. USA 90:4181–4185.
13. Baron, C., E. Westhof, A. Bck, and R. Giege. 1993. Solution structure of selenocysteine-inserting tRNASEC from Escherichia coli. J. Mol. Biol. 231:274–292.
14. Belaich, A., and J. P. Belaich. 1976. Microcalorimetric study of the anaerobic growth of Escherichia coli: growth thermograms in a synthetic medium. J. Bacteriol. 125:14–18.
15. Birkmann, A., and A. Bck. 1989. Characterisation of a cis regulatory DNA element necessary for formate induction of the formate dehydrogenase gene (fdhF) of Escherichia coli. Mol. Microbiol. 3:187–195.
16. Birkmann, A., R. G. Sawers, and A. Bck. 1987. Involvement of the ntrA gene product in the anaerobic metabolism of Escherichia coli. Mol. Gen. Genet. 210:535–542.
17. Birkmann, A., F. Zinoni, G. Sawers, and A. Bck. 1987. Factors affecting transcriptional regulation of the formate-hydrogen-lyase pathway of Escherichia coli. Arch. Microbiol. 148:44–51.
18. Blackwood, A. C., A. C. Neish, and G. A. Ledingham. 1956. Dissimilation of glucose at controlled pH values by pigmented and non-pigmented strains of Escherichia coli. J. Bacteriol. 72:497–499.
19. Blomqvist, K., M. Nokkola, P. Lehtovaara, M.-J. Suihko, U. Airaksinen, K. B. Strby, J. K. C. Knowles, and M. E. Penttil. 1993. Characterization of the genes of the 2,3-butanediol operons from Klebsiella terrigena and Enterobacter aerogenes. J. Bacteriol. 175:1392–1404.
20. Bck, A., K. Forchhammer, J. Heider, W. Leinfelder, G. Sawers, B. Veprek, and F. Zinoni. 1991. Selenocysteine: the 21st amino acid. Mol. Microbiol. 5:515–520.
21. Bhm, R., M. Sauter, and A. Bck. 1990. Nucleotide sequence and expression of an operon in Escherichia coli coding for formate hydrogenlyase components. Mol. Microbiol. 4:231–243.
22. Bokranz, M., M. Gutmann, C. Krtner, E. Kojro, F. Fahrenholz, F. Lauterbach, and A. Krger. 1991. Cloning and nucleotide sequence of the structural genes encoding the formate dehydrogenase of Wolinella succinogenes. Arch. Microbiol. 156:119–128.
23. Breed, R. S., and E. G. D. Murray. 1957. Family IV. Enterobacteriaceae, p. 333, 341–347, 350–351. In R. S. Breed, E. G. D. Murray, and N. R. Smith (ed.), Bergey’s Manual of Determinative Bacteriology. The Williams and Wilkins Co., Baltimore.
24. Brenner, D. J. 1984. Facultatively anaerobic gram-negative rods, p. 408–516. In N. R. Krieg and J. G. Holt (ed.), Bergey’s Manual of Systematic Bacteriology, vol. 1. The Williams and Wilkins Co., Baltimore.
25. Brenner, D. J. 1992. Introduction to the Enterobacteriaceae, p. 2673–2695. In A. Balows, H. G. Trper, M. Dworkin, W. Harder, and K.-H. Schleifer (ed.), The Prokaryotes. Springer-Verlag, New York.
26. Brown, T. D. K., M. C. Jones-Mortimer, and H. L. Kornberg. 1977. The enzymic interconversion of acetate and acetyl-coenzyme A in Escherichia coli. J. Gen. Microbiol. 102:327–336.
27. Brown, T. D. K., C. R. S. Pereira, and F. C. Strmer. 1972. Studies of the acetate kinase-phosphotransacetylase and the butanediol-forming systems in Aerobacter aerogenes. J. Bacteriol. 112:1106–1111.
28. Burk, R. F. 1991. Molecular biology of selenium with implications for its metabolism. FASEB J. 5:2274–2279.
29. Campbell, H. D., B. L. Rogers, and I. G. Young. 1984. Nucleotide sequence of the respiratory D-lactate dehydrogenase gene of Escherichia coli. Eur. J. Biochem. 144:367–373.
30. Casse, F., M.-C. Pascal, M. Chippaux, and J. Ratouchniak. 1976. Genetic analysis of mutants from Escherichia coli K12 unable to grow anaerobically without exogenous acceptor. Mol. Gen. Genet. 148:337–340.
31. Chen, Y.-M., and E. C. C. Lin. 1991. Regulation of the adhE gene, which encodes ethanol dehydrogenase in Escherichia coli. J. Bacteriol. 173:8009–8013.
32. Chevalier, M., E. C. C. Lin, and R. L. Levine. 1990. Hydrogen peroxide mediates the oxidative inactivation of enzymes following the switch from anaerobic to aerobic metabolism in Klebsiella pneumoniae. J. Biol. Chem. 265:42–46.
33. Clark, D., and J. E. Cronan, Jr. 1980. Escherichia coli mutants with altered control of alcohol dehydrogenase and nitrate reductase. J. Bacteriol. 141:177–183.
34. Clark, D. P. 1989. The fermentation pathways of Escherichia coli. FEMS Microbiol. Rev. 63:223–234.
35. Clark, D. P., and J. E. Cronan, Jr. 1980. Acetaldehyde coenzyme A dehydrogenase of Escherichia coli. J. Bacteriol. 144:179–184.
36. Clark, D. P., P. R. Cunningham, S. G. Reams, F. Mat-Jan, R. Mohammedkhani, and C. R. Williams. 1988. Mutants of Escherichia coli defective in acid fermentation. Appl. Biochem. Biotechnol. 17:163–173.
37. Conway, T., G. W. Sewell, Y. A. Osman, and L. O. Ingram. 1987. Cloning and sequencing of the alcohol dehydrogenase II gene from Zymomonas mobilis. J. Bacteriol. 169:2591–2597.
38. Cox, J. C., E. S. Edwards, and J. A. DeMoss. 1981. Resolution of distinct selenium-containing formate dehydrogenases from Escherichia coli. J. Bacteriol. 145:1317–1324.
39. Cunningham, P. R., and D. P. Clark. 1986. The use of suicide substrates to select mutants of Escherichia coli lacking enzymes of alcohol fermentation. Mol. Gen. Genet. 205:487–493.
40. Daniel, R., and G. Gottschalk. 1992. Growth temperature-dependent activity of glycerol dehydratase in Escherichia coli expressing the Citrobacter freundii dha regulon. FEMS Microbiol. Lett. 100:281–286.
41. Dawes, E. A., and S. M. Foster. 1956. The formation of ethanol in Escherichia coli. Biochim. Biophys. Acta 22:253–265.
42. Dekker, E. E., and R. R. Swain. 1968. Formation of D-1-amino-2-propanol by a highly purified enzyme from Escherichia coli. Biochim. Biophys. Acta 158:306–307.
43. Dong, J. M., J. S. Taylor, D. J. Latour, S. Iuchi, and E. C. C. Lin. 1993. Three overlapping lct genes involved in l-lactate utilization by Escherichia coli. J. Bacteriol. 175:6671–6678.
44. Dross, F., V. Geisler, R. Lenger, F. Theis, T. Kraft, F. Fahrenholz, E. Kojro, A. Duchene, D. Tripier, K. Juvenal, and A. Krger. 1992. The quinone-reactive Ni/Fe-hydrogenase of Wolinella succinogenes. Eur. J. Biochem. 206:93–102.
45. Eddy, B. P. 1961. The Voges-Proskauer reaction and its significance: a review. J. Appl. Bacteriol. 24:27–41.
46. Ehrenreich, A., K. Forchhammer, P. Tormay, B. Veprek, and A. Bck. 1992. Selenoprotein synthesis in E. coli: purification and characterisation of the enzyme catalysing selenium activation. Eur. J. Biochem. 206:767–773.
47. Engelhardt, H., K. Forchhammer, S. Mller, K. N. Goldie, and A. Bck. 1992. Structure of selenocysteine synthase from Escherichia coli and location of tRNA in the seryl-tRNASEC-enzyme complex. Mol. Microbiol. 6:3461–3467.
48. Enoch, H. G., and R. L. Lester. 1975. The purification and properties of formate dehydrogenase and nitrate reductase from Escherichia coli. J. Biol. Chem. 250:6693–6705.
49. Forage, R. G., and M. A. Foster. 1979. Resolution of the coenzyme B-12-dependent dehydratases of Klebsiella sp. and Citrobacter freundii. Biochim. Biophys. Acta 569:249–258.
50. Forage, R. G., and M. A. Foster. 1982. Glycerol fermentation in Klebsiella pneumoniae: functions of the coenzyme B12-dependent glycerol and diol dehydratases. J. Bacteriol. 149:413–419.
51. Forage, R. G., and E. C. C. Lin. 1982. dha system mediating aerobic and anaerobic dissimilation of glycerol in Klebsiella pneumoniae NCIB 418. J. Bacteriol. 151:591–599.
52. Forchhammer, K., and A. Bck. 1991. Selenocysteine synthase from Escherichia coli: analysis of the reaction sequence. J. Biol. Chem. 266:6324–6328.
53. Forchhammer, K., W. Leinfelder, and A. Bck. 1989. Identification of a novel translation factor necessary for the incorporation of selenocysteine into protein. Nature (London) 342:453–456.
54. Forchhammer, K., W. Leinfelder, K. Boesmiller, B. Veprek, and A. Bck. 1991. Selenocysteine synthase from Escherichia coli: nucleotide sequence of the gene (selA) and purification of the protein. J. Biol. Chem. 266:6318–6323.
55. Frster, C., G. Ott, K. Forchhammer, and M. Sprinzl. 1990. Interaction of a selenocysteine-incorporating tRNA with elongation factor Tu from E. coli. Nucleic Acids Res. 18:487–491.
56. Fox, D. K., N. D. Meadow, and S. Roseman. 1986. Phosphate transfer between acetate kinase and enzyme I of the bacterial phosphotransferase system. J. Biol. Chem. 261:13498–13503.
57. Fox, D. K., and S. Roseman. 1986. Isolation and characterization of homogeneous acetate kinase from Salmonella typhimurium and Escherichia coli. J. Biol. Chem. 261:13487–13497.
58. Francis, K., P. Patel, J. C. Wendt, and K. T. Shanmugam. 1990. Purification and characterization of two forms of hydrogenase isoenzyme 1 from Escherichia coli. J. Bacteriol. 172:5750–5757.
59. Friedrich, B., and E. Schwartz. 1993. Molecular biology of hydrogen utilization in aerobic chemolithotrophs. Annu. Rev. Microbiol. 47:351–383.
60. Futai, M., and H. Kimura. 1977. Inducible membrane bound D-lactate dehydrogenase from Escherichia coli. Purification and properties. J. Biol. Chem. 252:5820–5827.
61. Garvie, E. I. 1980. Bacterial lactate dehydrogenases. Microbiol. Rev. 44:106–139.
62. Gest, H., and H. D. Peck, Jr. 1955. A study of the hydrogenlyase reaction with systems derived from normal and anaerogenic coli-aerogenes bacteria. J. Bacteriol. 70:326–334.
63. Giordano, G., C.-L. Medani, M.-A. Mandrand-Berthelot, and D. H. Boxer. 1983. Formate dehydrogenase from Escherichia coli. FEMS Microbiol. Lett. 17:171–177.
64. Godtfredsen, S. E., H. Lorck, and P. Sigsgaard. 1983. On the occurrence of α-acetolactate decarboxylases among microorganisms. Carlsberg Res. Commun. 48:239–247.
65. Gollin, D. J., L. E. Mortenson, and R. L. Robson. 1992. Carboxyl-terminal processing may be essential for production of active NiFe hydrogenase in Azotobacter vinelandii. FEBS Lett. 309:371–375.
66. Goodlove, P. E., P. R. Cunningham, J. Parker, and D. P. Clark. 1989. Cloning and sequence analysis of the fermentative alcohol-dehydrogenase-encoding gene of Escherichia coli. Gene 85:209–214.
67. Graham, A. 1981. The organisation of hydrogenase in the cytoplasmic membrane of Escherichia coli. Biochem. J. 197:283–291.
68. Grimont, F., and P. A. D. Grimont. 1992. The genus Enterobacter, p. 2797–2815. In A. Balows, H. G. Trper, M. Dworkin, W. Harder, and K.-H. Schleifer (ed.), The Prokaryotes. Springer-Verlag, New York.
69. Grimont, F., P. A. D. Grimont, and C. Richard. 1992. The genus Klebsiella, p. 2775–2796. In A. Balows, H. G. Trper, M. Dworkin, W. Harder, and K.-H. Schleifer (ed.), The Prokaryotes. Springer-Verlag, New York.
70. Guest, J. R. 1979. Anaerobic fermentation of E. coli K12 with fumarate as terminal electron acceptor. Genetic studies with menaquinone and fluoroacetate-resistant mutants. J. Gen. Microbiol. 115:259–271.
71. Guest, J. R., S. J. Angier, and G. C. Russell. 1989. Structure, expression, and protein engineering of the pyruvate dehyrogenase complex of Escherichia coli. Ann. N.Y. Acad. Sci. 573:76–99.
72. Gupta, S., and D. P. Clark. 1989. Escherichia coli derivatives lacking both alcohol dehydrogenase and phosphotransacetylase grow anaerobically by lactate fermentation. J. Bacteriol. 171:3650–3655.
73. Hansen, R. G., and U. Henning. 1966. Regulation of pyruvate dehydrogenase activity in Escherichia coli K12. Biochim. Biophys. Acta 122:355–358.
74. Harden, A., and G. S. Walpole. 1906. Chemical action of Bacillus lactis aerogenes (Escherich) on glucose and mannitol: production of 2,3-butyleneglycol and acetylmethylcarbinol. Proc. R. Soc. London Ser. B 77:399–404.
75. Hausinger, R. P. 1987. Nickel utilization by microorganisms. Microbiol. Rev. 51:22–42.
76. Heider, J., C. Baron, and A. Bck. 1992. Coding from a distance: dissection of the mRNA determinants required for the incorporation of selenocysteine into protein. EMBO J. 11:3759–3766.
77. Heider, J., and A. Bck. 1992. Targeted insertion of selenocysteine into the α subunit of formate dehydrogenase from Methanobacterium formicicum. J. Bacteriol. 174:659–663.
78. Heider, J., and A. Bck. 1993. Selenium metabolism in micro-organisms. Adv. Microb. Physiol. 35:71–109.
79. Hickman, J., and G. Ashwell. 1960. Uronic acid metabolism in bacteria. II. Purification and properties of D-altronic and D-mannonic acid dehydrogenases in Escherichia coli. J. Biol. Chem. 235:1566–1570.
80. Higgins, C. F., M. P. Gallacher, S. C. Hyde, M. L. Mimmack, and S. R. Pearce. 1990. Periplasmic binding protein-dependent transport systems: the membrane associated components. Philos. Trans. R. Soc. London 326:353–365.
81. Hohn-Bentz, H., and F. Radler. 1978. Bacterial 2,3-butanediol dehydrogenases. Arch. Mikrobiol. 116:197–203.
82. Holtzclaw, W. D., and L. F. Chapman. 1975. Degradative acetolactate synthase of Bacillus subtilis: purification and properties. J. Bacteriol. 121:917–922.
83. Hopper, S., M. Babst, V. Schlensog, H.-M. Fischer, H. Hennecke, and A. Bck. 1994. Regulated expression in vitro of genes coding for formate hydronelyase components of Escherichia coli. J. Biol. Chem. 269:19597–19604.
84. Hormaeche, E., and P. R. Edwards. 1960. A proposed genus Enterobacter. Int. Bull. Bacteriol. Nomencl. Taxon. 10:71–74.
85. Huseby, N.-E., T. B. Christensen, B. J. Olsen, and F. C. Strmer. 1971. The pH 6 acetolactate-forming enzyme from Acetobacter aerogenes. Subunit structure. Eur. J. Biochem. 20:209–214.
86. Jacobi, A., R. Rossmann, and A. Bck. 1992. The hyp operon gene products are required for the maturation of catalytically active hydrogenase isoenzymes in Escherichia coli. Arch. Microbiol. 158:444–451.
87. Jamieson, D. J., R. G. Sawers, P. A. Rugman, D. H. Boxer, and C. F. Higgins. 1986. Effects of anaerobic regulatory mutations and catabolite repression on regulation of hydrogen metabolism and hydrogenase isoenzyme composition in Salmonella typhimurium. J. Bacteriol. 168:405–411.
88. Janson, C. A., and W. W. Cleland. 1974. The inhibition of acetate, pyruvate, and 3-phosphoglycerate kinases by chromium adenosine triphosphate. J. Biol. Chem. 249:2567–2571.
89. Jin, R., and E. C. C. Lin. 1984. An inducible phosphoenolpyruvate:dihydroxyacetone phosphotransferase system in Escherichia coli. J. Gen. Microbiol. 130:83–88.
90. Jin, R. Z., J. C.-T. Tang, and E. C. C. Lin. 1983. Experimental evolution of a novel pathway for glycerol dissimilation in Escherichia coli. J. Mol. Evol. 19:429–436.
91. Johansen, L., K. Bryn, and F. C. Strmer. 1975. Physiological and biochemical role of the butanediol pathway in Aerobacter (Enterobacter) aerogenes. J. Bacteriol. 123:1124–1130.
92. Johnson, E. A., S. K. Burke, R. G. Forage, and E. C. C. Lin. 1984. Purification and properties of dihydroxyacetone kinase from Klebsiella pneumoniae. J. Bacteriol. 160:55–60.
93. Johnson, E. A., R. L. Levine, and E. C. C. Lin. 1985. Inactivation of glycerol dehydrogenase of Klebsiella pneumoniae and the role of divalent cations. J. Bacteriol. 164:479–483.
94. Jones, R. W. 1980. Proton translocation by the membrane-bound formate dehydrogenase of Escherichia coli. FEMS Microbiol. Lett. 8:167–171.
95. Kaiser, M., and G. Sawers. 1994. Pyruvate formate-lyase is not essential for nitrate respiration by Escherichia coli. FEMS Microbiol. Lett. 117:163–168.
96. Karube, I., M. Tomiyama, and A. Kikuchi. 1984. Molecular cloning and physical mapping of the hyd gene of Escherichia coli K-12. FEMS. Microbiol. Lett. 25:165–168.
97. Kelley, J. J., and E. E. Dekker. 1985. Identity of Escherichia coli d-1-amino-2-propanol:NAD+ oxidoreductase with E. coli glycerol dehydrogenase but not with Neisseria gonorrhoeae 1,2-propanediol:NAD+ oxidoreductase. J. Bacteriol. 162:170–175.
98. Kessler, D., W. Herth, and J. Knappe. 1992. Ultrastructure and pyruvate-formate-lyase quenching property of the multienzymic AdhE protein of Escherichia coli. J. Biol. Chem. 267:18073–18079.
99. Kessler, D., I. Leibrecht, and J. Knappe. 1991. Pyruvate-formate-lyase-deactivase and acetyl-CoA reductase activities of Escherichia coli reside on a polymeric protein particle encoded by adhE. FEBS Lett. 281:59–63.
100. Kim, I. Y., Z. Veres, and T. C. Stadtman. 1992. Biochemical analysis of Escherichia coli selenophosphate synthetase mutants: lysine-20 is essential for catalytic activity and cysteine-17/19 for 8-azido-ATP derivatization. J. Biol. Chem. 267:19650–19654.
101. Knappe, J., and G. Sawers. 1990. A radical-chemical route to acetyl-CoA: the anaerobically induced pyruvate formate-lyase system of Escherichia coli. FEMS Microbiol. Rev. 75:383–398.
102. Krasna, A. I. 1984. Mutants of Escherichia coli with altered hydrogenase activity. J. Gen. Microbiol. 130:779–787.
103. Kukuruzinska, M. A., B. W. Turner, G. K. Ackers, and S. Roseman. 1984. Subunit association of enzyme I of the Salmonella typhimurium phosphoenolpyruvate:glycose phosphotransferase system: temperature dependence and thermodynamic properties. J. Biol. Chem. 259:11679–11681.
104. Kwan, H. S., H. W. Chui, and K. K. Wong. 1988. ack::Mud1–8 (Aprlac) operon fusions of Salmonella typhimurium LT2. Mol. Gen. Genet. 211:183–185.
105. Langley, D., and J. R. Guest. 1977. Biochemical genetics of the α-keto acid dehydrogenase complexes of Escherichia coli K12: isolation and biochemical properties of deletion mutants. J. Gen. Microbiol. 99:263–276.
106. Lee, J. H., P. Patel, P. Sankar, and K. T. Shanmugam. 1985. Isolation and characterization of mutant strains of Escherichia coli altered in hydrogen metabolism. J. Bacteriol. 162:344–352.
107. Leinfelder, W., K. Forchhammer, B. Veprek, E. Zehelein, and A. Bck. 1990. In vitro synthesis of selenocysteinyl-tRNAUCA from seryl-tRNAUCA: involvement and characterization of the selD gene product. Proc. Natl. Acad. Sci. USA 87:543–547.
108. Leinfelder, W., K. Forchhammer, F. Zinoni, G. Sawers, M.-A. Mandrand-Berthelot, and A. Bck. 1988. Escherichia coli genes whose products are involved in selenium metabolism. J. Bacteriol. 170:540–546.
109. Leinfelder, W., E. Zehelein, M.-A. Mandrand-Berthelot, and A. Bck. 1988. Gene for a novel tRNA species that accepts L-serine and cotranslationally inserts selenocysteine. Nature (London) 331:723–725.
110. Leonardo, M. R., P. R. Cunningham, and D. P. Clark. 1993. Anaerobic regulation of the adhE gene, encoding the fermentative alcohol dehydrogenase of Escherichia coli. J. Bacteriol. 175:870–878.
111. LeVine, S., F. Ardeshir, and G. F.-L. Ames. 1980. Isolation and characterization of acetate kinase and phosphotransacetylase mutants of Escherichia coli and Salmonella typhimurium. J. Bacteriol. 143:1081–1085.
112. Li, J., and V. Stewart. 1992. Localization of upstream sequence elements required for nitrate and anaerobic induction of fdn (formate dehydrogenase-N) operon expression in Escherichia coli K-12. J. Bacteriol. 174:4935–4942.
113. Lin, E. C. C., A. Levin, and B. Magasanik. 1960. The effect of aerobic metabolism on the inducible glycerol dehydrogenase of Aerobacter aerogenes. J. Biol. Chem. 235:1824–1829.
114. Lken, J. P., and F. C. Strmer. 1970. Acetolactate decarboxylase from Aerobacter aerogenes. Purification and properties. Eur. J. Biochem. 14:133–137.
115. Lorowitz, W., and D. Clark. 1982. Escherichia coli mutants with a temperature-sensitive alcohol dehydrogenase. J. Bacteriol. 152:935–938.
116. Lutz, S., R. Bhm, A. Beier, and A. Bck. 1990. Characterization of divergent NtrA-dependent promoters in the anaerobically expressed gene cluster coding for hydrogenase 3 components of Escherichia coli. Mol. Microbiol. 4:13–20.
117. Lutz, S., A. Jacobi, V. Schlensog, R. Bhm, G. Sawers, and A. Bck. 1991. Molecular characterization of an operon (hyp) necessary for the activity of the three hydrogenase isoenzymes in Escherichia coli. Mol. Microbiol. 5:123–135.
118. Magasanik, B., M. S. Brooke, and D. Karibian. 1953. Metabolic pathways of glycerol dissimilation. J. Bacteriol. 66:611–619.
119. Magee, R. J., and N. Kosaric. 1987. The microbial production of 2,3-butanediol. Adv. Appl. Microbiol. 32:89–161.
120. Maier, T., A. Jacobi, M. Sauter, and A. Bck. 1993. The product of the hypB gene, which is required for nickel incorporation into hydrogenases, is a novel guanine nucleotide-binding protein. J. Bacteriol. 175:630–635.
121. Maloy, S. R., and W. D. Nunn. 1982. Genetic regulation of the glyoxylate shunt in Escherichia coli K-12. J. Bacteriol. 149:173–180.
122. Malthe-Srenssen, D., and F. C. Strmer. 1970. The pH 6 acetolactate-forming enzyme from Serratia marcescens. Purification and properties. Eur. J. Biochem. 14:127–132.
123. Matayoshi, S., H. Oda, and G. Sarwar. 1989. Relationship between the production of spirosomes and anaerobic glycolysis activity in Escherichia coli. J. Gen. Microbiol. 135:525–529.
124. Mat-Jan, F., K. Y. Alam, and D. P. Clark. 1989. Mutants of Escherichia coli deficient in the fermentative lactate dehydrogenase. J. Bacteriol. 171:342–348.
125. Matsuyama, A., H. Yamamoto, and E. Nakano. 1989. Cloning, expression, and nucleotide sequence of the Escherichia coli K-12 ackA gene. J. Bacteriol. 171:577–580.
126. McGregor, W. G., J. Phillips, and C. H. Suelter. 1974. Purification and kinetic characterization of a monovalent cation-activated glycerol dehydrogenase from Aerobacter aerogenes. J. Biol. Chem. 249:3132–3139.
127. Menon, N. K., J. Robbins, H. D. Peck, C. Y. Chatelus, E.-S. Choi, and A. E. Przybyla. 1990. Cloning and sequencing of a putative Escherichia coli (NiFe) hydrogenase-1 operon containing six open reading frames. J. Bacteriol. 172:1969–1977.
128. Menon, N. K., J. Robbins, J. C. Wendt, K. T. Shanmugam, and A. E. Przybyla. 1991. Mutational analysis and characterization of the Escherichia coli hya operon, which encodes (NiFe) hydrogenase 1. J. Bacteriol. 173:4851–4861.
129. Michels, P. A. M., J. P. J. Michels, J. Boonstra, and W. N. Konings. 1979. Generation of an electrochemical proton gradient in bacteria by the excretion of metabolic end products. FEMS Microbiol. Lett. 5:357–364.
130. Navarro, C., L.-F. Wu, and M.-A. Mandrand-Berthelot. 1993. The nik operon of Escherichia coli encodes a periplasmic binding-protein-dependent transport sytem for nickel. Mol. Microbiol. 9:1181–1191.
131. Nishimura, Y., I. K. P. Tan, Y. Ohgami, K. Kohgami, and T. Kamihara. 1983. Induction of membrane-bound L-lactate dehydrogenase in Escherichia coli under conditions of nitrate respiration, fumarate reduction and trimethylamine-N-oxide reduction. FEMS Microbiol. Lett. 17:283–286.
132. Novotny, M. J., J. Reizer, F. Esch, and M. H. Saier, Jr. 1984. Purification and properties of d-mannitol-1-phosphate dehydrogenase and d-glucitol-6-phosphate dehydrogenase from Escherichia coli. J. Bacteriol. 159:986–990.
133. Ogino, T., Y. Arata, and S. Fujiwara. 1980. Proton correlation nuclear magnetic resonance study of metabolic regulations and pyruvate transport in anaerobic Escherichia coli cells. Biochemistry 19:3684–3691.
134. Ogino, T., Y. Arata, S. Fujiwara, H. Shoun, and T. Beppu. 1978. Proton correlation nuclear magnetic resonance study of anaerobic metabolism of Escherichia coli. Biochemistry 17:4742–4745.
135. Ohshiro, T., K. Aisaka, and T. Uwajima. 1989. Purification and characterization of α-acetolactate decarboxylase from Brevibacterium acetylicum. Agric. Biol. Chem. 53:1913–1918.
136. Park, M. H., B. B. Wong, and J. E. Lusk. 1976. Mutants in three genes affecting transport of magnesium in Escherichia coli: physiology and genetics. J. Bacteriol. 126:1096–1103.
137. Pascal, M. C., M. Chippaux, A. Abou-Jaoude, H. P. Blaschkowski, and J. Knappe. 1981. Mutants of Escherichia coli K12 with defects in anaerobic pyruvate metabolism. J. Gen. Microbiol. 124:35–42.
138. Pecher, A., F. Zinoni, and A. Bck. 1985. The seleno-polypeptide of formic dehydrogenase (formate hydrogen-lyase linked) from Escherichia coli: genetic analysis. Arch. Microbiol. 141:359–363.
139. Pecher, A., F. Zinoni, C. Jatisatienr, R. Wirth, H. Hennecke, and A. Bck. 1983. On the redox control of synthesis of anaerobically induced enzymes in enterobacteriaceae. Arch. Microbiol. 136:131–136.
140. Peck, H. D., and H. Gest. 1957. Formic dehydrogenase and the hydrogenlyase enzyme complex in coli-aerogenes bacteria. J. Bacteriol. 73:706–721.
141. Pinsent, J. 1954. The need for selenite and molybdate in the formation of formic dehydrogenase by members of the coli-aerogenes. Biochem. J. 57:10–16.
142. Przybyla, A. E., J. Robbins, N. Menon, and H. D. Peck. 1992. Structure-function relationships among nickel-containing hydrogenases. FEMS Microbiol. Rev. 88:109–136.
143. Rasmussen, A. M., R. M. Gibson, S. E. Godtfredsen, and M. Ottesen. 1985. Purification of α-acetolactate decarboxylase from Lactobacillus casei DSM 2547. Carlsberg Res. Commun. 50:73–82.
144. Reeve, J. N., and G. S. Beckler. 1990. Conservation of primary structure in prokaryotic hydrogenases. FEMS Microbiol. Rev. 87:419–424.
145. Renna, M. C., N. Najimudin, L. R. Winik, and S. A. Zahler. 1993. Regulation of the Bacillus subtilis alsS, alsD, and alsR genes involved in the post-exponential-phase production of acetoin. J. Bacteriol. 175:3863–3875.
146. Reynolds, H., and C. H. Werkman. 1937. The intermediate dissimilation of glucose by Aerobacter indologenes. J. Bacteriol. 33:603–614.
147. Rossmann, R., M. Sauter, F. Lottspeich, and A. Bck. 1994. Maturation of the large subunit (HycE) of hydrogenase 3 of Escherichia coli requires nickel incorporation followed by C-terminal processing at Arg537. Eur. J. Biochem. 220:377–384.
148. Rossmann, R., G. Sawers, and A. Bck. 1991. Mechanism of regulation of the formate-hydrogenlyase pathway by oxygen, nitrate, and pH: definition of the formate regulon. Mol. Microbiol. 5:2807–2814.
149. Ruch, F. E., J. Lengler, and E. C. C. Lin. 1974. Regulation of glycerol catabolism in Klebsiella aerogenes. J. Bacteriol. 119:50–56.
150. Ruch, F. E., and E. C. C. Lin. 1975. Independent constitutive expression of the aerobic and anaerobic pathways of glycerol catabolism in Klebsiella aerogenes. J. Bacteriol. 124:348–352.
151. Ruch, F. E., E. C. C. Lin, J. D. Kowit, C.-T. Tang, and A. L. Goldberg. 1980. In vivo inactivation of glycerol dehydrogenase in Klebsiella aerogenes: properties of active and inactivated proteins. J. Bacteriol. 141:1077–1085.
152. Rudolph, F. B., D. L. Purich, and H. J. Fromm. 1968. Coenzyme A-linked aldehyde dehydrogenase from Escherichia coli: partial purification, properties, and kinetic studies of the enzyme. J. Biol. Chem. 243:5539–5545.
153. Ruiz-Herrera, J., and A. Alvarez. 1972. A physiological study of formate dehydrogenase, formate oxidase and hydrogenlyase from Escherichia coli K-12. Antonie van Leeuwenhoek 38:479–491.
154. Rush, D., D. Kiribian, M. L. Karnovsky, and B. Magasanik. 1957. Pathways of glycerol dissimilation in two strains of Aerobacter aerogenes: enzymatic and tracer studies. J. Biol. Chem. 226:891–899.
155. Sanwal, B. D. 1970. Allosteric controls of amphibolic pathways in bacteria. Bacteriol. Rev. 34:20–39.
156. Sauter, M., R. Bhm, and A. Bck. 1992. Mutational analysis of the operon (hyc) determining hydrogenase 3 formation in Escherichia coli. Mol. Microbiol. 6:1523–1532.
157. Sawers, G. 1993. Specific transcriptional requirements for positive regulation of the anaerobically inducible pfl operon by ArcA and FNR. Mol. Microbiol. 10:737–747.
158. Sawers, G. 1994. The hydrogenases and formate dehydrogenases of Escherichia coli. Antonie van Leuwenhoek J. Microbiol. 66:57–88.
159. Sawers, G., and A. Bck. 1988. Anaerobic regulation of pyruvate formate-lyase from Escherichia coli K-12. J. Bacteriol. 170:5330–5336.
160. Sawers, G., and A. Bck. 1989. Novel transcriptional control of the pyruvate formate-lyase gene: upstream regulatory sequences and multiple promoters regulate anaerobic expression. J. Bacteriol. 171:2485–2498.
161. Sawers, G., J. Heider, E. Zehelein, and A. Bck. 1991. Expression and operon structure of the sel genes of Escherichia coli and identification of a third selenium-containing formate dehydrogenase isoenzyme. J. Bacteriol. 173:4983–4993.
162. Sawers, G., and B. Suppmann. 1992. Anaerobic induction of pyruvate formate-lyase gene expression is mediated by the ArcA and FNR proteins. J. Bacteriol. 174:3474–3478.
163. Sawers, G., A. F. V. Wagner, and A. Bck. 1989. Transcription initiation at multiple promoters of the pfl gene by Eσ70-dependent transcription in vitro and heterologous expression in Pseudomonas putida in vivo. J. Bacteriol. 171:4930–4937.
164. Sawers, R. G., S. P. Ballantine, and D. H. Boxer. 1985. Differential expression of hydrogenase isoenzymes in Escherichia coli K-12: evidence for a third isoenzyme. J. Bacteriol. 164:1324–1331.
165. Sawers, R. G., and D. H. Boxer. 1986. Purification and properties of membrane-bound hydrogenase isoenzyme 1 from anaerobically grown Escherichia coli K12. Eur. J. Biochem. 156:265–275.
166. Sawers, R. G., D. J. Jamieson, C. F. Higgins, and D. H. Boxer. 1986. Characterization and physiological roles of membrane-bound hydrogenase isoenzymes from Salmonella typhimurium. J. Bacteriol. 168:398–404.
167. Schlensog, V., A. Birkmann, and A. Bck. 1989. Mutations in trans which affect the anaerobic expression of a formate dehydrogenase (fdhF) structural gene. Arch. Microbiol. 152:83–89.
168. Schlensog, V., and A. Bck. 1990. Identification and sequence analysis of the gene encoding the transcriptional activator of the formate hydrogenlyase system of Escherichia coli. Mol. Microbiol. 4:1319–1327.
169. Schlensog, V., S. Lutz, and A. Bck. 1994. Purification and DNA-binding properties of FHLA, the transcriptional activator of the formate hydrogenlyase system from E. coli. J. Biol. Chem. 269:19590–19596.
170. Schmitt, B. 1975. Aldehyde dehydrogenase activity of a complex particle from E. coli. Biochimie 57:1001–1004.
171. Shimizu, M., T. Suzuki, K.-Y. Kameda, and Y. Abiko. 1969. Phosphotransacetylase of Escherichia coli B, purification and properties. Biochim. Biophys. Acta 191:550–558.
172. Shuber, A. P., E. C. Orr, M. A. Recny, P. F. Schendel, H. D. May, N. L. Schauer, and J. G. Ferry. 1986. Cloning, expression, and nucleotide sequence of the formate dehydrogenase genes from Methanobacterium formicicum. J. Biol. Chem. 261:12943–12947.
173. Sirko, A., E. Zehelein, M. Freundlich, and G. Sawers. 1993. Integration host factor is required for anaerobic pyruvate induction of pfl operon expression in Escherichia coli. J. Bacteriol. 175:5769–5777.
174. Sokatch, J. R. 1969. Bacterial Physiology and Metabolism. Academic Press, Ltd., London.
175. Sprenger, G. A., B. A. Hammer, E. A. Johnson, and E. C. C. Lin. 1989. Anaerobic growth of Escherichia coli on glycerol by importing genes of the dha regulon from Klebsiella pneumoniae. J. Gen. Microbiol. 135:1255–1262.
176. St. Martin, E. J., W. B. Freedberg, and E. C. C. Lin. 1977. Kinase replacement by a dehydrogenase for Escherichia coli glycerol utilization. J. Bacteriol. 131:1026–1028.
177. Stadtman, T. C. 1990. Selenium biochemistry. Annu. Rev. Biochem. 59:111–127.
178. Stadtman, T. C. 1991. Biosynthesis and function of selenocysteine-containing enzymes. J. Biol. Chem. 59:16257–16260.
179. Stadtman, T. C., J. N. Davis, W.-M. Ching, F. Zinoni, and A. Bck. 1991. Amino acid sequence analysis of Escherichia coli formate dehydrogenase (FDHH) confirms that TGA in the gene encodes selenocysteine in the gene product. Biofactors 3:21–27.
180. Stephenson, M., and L. H. Stickland. 1931. Hydrogenase: a bacterial enzyme activating molecular hydrogen. I. The properties of the enzyme. Biochem. J. 25:205–214.
181. Stephenson, M., and L. H. Stickland. 1932. Hydrogenlyases. Bacterial enzymes liberating molecular hydrogen. Biochem. J. 26:712–724.
182. Stoker, K., L. F. Oltmann, and A. H. Stouthammer. 1989. Randomly induced Escherichia coli K-12 Tn5 insertion mutants defective in hydrogenase activity. J. Bacteriol. 171:831–836.
183. Stokes, J. L. 1949. Fermentation of glucose by suspensions of Escherichia coli. J. Bacteriol. 57:147–158.
184. Strmer, F. C. 1968. Evidence for induction of the 2,3-butanediol-forming enzymes in Aerobacter aerogenes. FEBS Lett. 2:36–28.
185. Strmer, F. C. 1968. The pH 6 acetolactate-forming enzyme from Aerobacter aerogenes. I. Kinetic studies. J. Biol. Chem. 243:3735–3739.
186. Strmer, F. C. 1975. 2,3-butanediol biosynthetic system in Aerobacter aerogenes. Methods Enzymol. 41B:518–533.
187. Strmer, F. C. 1977. Evidence for regulation of Aerobacter aerogenes pH 6 acatolactate-forming enzyme by acetate ions. Biochem. Biophys. Res. Commun. 74:898–902.
188. Stroinski, A., J. Pawelkiewicz, and B. C. Johnson. 1974. Allosteric interactions in glycerol dehydratase: purification of enzyme and effects of positive and negative cooperativity for glycerol. Arch. Biochem. Biophys. 162:321–330.
189. Suppmann, B., and G. Sawers. 1994. Isolation and characterization of hypophosphite-resistant mutants of Escherichia coli: identification of the FocA protein, encoded by the pfl operon, as a putative formate transporter. Mol. Microbiol. 11:965–982.
190. Suzuki, T. 1969. Phosphotransacetylase of Escherichia coli B, activation by pyruvate and inhibition by NADH and certain nucleotides. Biochim. Biophys. Acta 191:559–569.
191. Svendsen, I., B. R. Jensen, and M. Ottesen. 1989. Complete amino acid sequence of α-acetolactate decarboxylase from Bacillus brevis. Carlsberg Res. Commun. 54:157–183.
192. Tang, C.-T., F. E. Ruch, Jr., and E. C. C. Lin. 1979. Purification and properties of a nicotinamide adenine dinucleotide-linked dehydrogenase that serves an Escherichia coli mutant for glycerol catabolism. J. Bacteriol. 140:182–187.
193. Tarmy, E. M., and N. O. Kaplan. 1965. Interacting binding sites of L-specific lactic dehydrogenase of Escherichia coli. Biochem. Biophys. Res. Commun. 21:379–383.
194. Tarmy, E. M., and N. O. Kaplan. 1968. Chemical characterization of D-lactate dehydrogenase from Escherichia coli B. J. Biol. Chem. 243:2579–2586.
195. Tarmy, E. M., and N. O. Kaplan. 1968. Kinetics of Escherichia coli B D-lactate dehydrogenase and evidence for pyruvate-controlled change in conformation. J. Biol. Chem. 243:2587–2596.
196. Ten Brink, B., and W. N. Konings. 1980. Generation of an electrochemical proton gradient by lactate efflux in membrane vesicles of Escherichia coli. Eur. J. Biochem. 111:59–66.
197. Thauer, R. K., K. Jungermann, and K. Decker. 1977. Energy conservation in chemotrophic anaerobic bacteria. Bacteriol. Rev. 41:100–180.
198. Tikka, J. 1935. ber den Mechanismus Glucosevergrung durch B. coli. Biochem. Z. 279:264–275.
199. Toraya, T., S. Honda, S. Kuno, and S. Fukui. 1978. Coenzyme B12-dependent diol dehydratase: regulation of apoenzyme synthesis in Klebsiella pneumoniae (Aerobacter aerogenes) ATCC8724. J. Bacteriol. 135:726–729.
200. Toraya, T., S. Kuno, and S. Fukui. 1980. Distribution of coenzyme B12-dependent diol dehydratase and glycerol dehydratase in selected genera of Enterobacteriaceae and Propionibacteriaceae. J. Bacteriol. 141:1439–1442.
201. Truniger, V., and W. Boos. 1994. Mapping and cloning of gldA, the structural gene of the Escherichia coli glycerol dehydrogenase. J. Bacteriol. 176:1796– 1800.
202. Varenne, S., F. Casse, M. Chippaux, and M. C. Pascal. 1975. A mutant of Escherichia coli deficient in pyruvate formate-lyase. Mol. Gen. Genet. 141:181–184.
203. Veres, Z., L. Tsai, T. D. Scholz, M. Politino, R. S. Balaban, and T. C. Stadtman. 1992. Synthesis of 5-methylaminomethyl-2-selenouridine in tRNAs: 31P NMR studies show how the labile selenium donor synthesized by the selD gene product contains selenium bonded to phosphorus. Proc. Natl. Acad. Sci. USA 89:2975–2979.
204. Vignais, P. M., and B. Toussaint. 1994. Molecular biology of membrane-bound H2 uptake hydrogenases. Arch. Microbiol. 161:1–10.
205. Wagner, A. F. V., M. Frey, F. A. Neugebauer, W. Schfer, and J. Knappe. 1992. The free radical in pyruvate formate-lyase is located on glycine-734. Proc. Natl. Acad. Sci. USA 89:996–1000.
206. Wagner, O. W., H. A. Lee, Jr., P. A. Frey, and R. H. Abeles. 1966. Studies on the mechanism of action of cobamide enzymes: chemical properties of the enzyme-coenzyme complex. J. Biol. Chem. 241:1751–1762.
207. Wanner, B. L., and M. R. Wilmes-Riesenberg. 1992. Involvement of phosphotransacetylase, acetate kinase, and acetyl phosphate synthesis in control of the phosphate regulon in Escherichia coli. J. Bacteriol. 174:2124–2130.
208. Weiss, H., T. Friedrich, G. Hofhaus, and D. Preis. 1991. The respiratory chain NADH dehydrogenase (complex I) of mitochondria. Eur. J. Biochem. 197:563–576.
209. Williamson, V. M., and C. E. Paquin. 1987. Homology of Saccharomyces cerevisiae ADH4 to an iron-activated alcohol dehydrogenase from Zymomonas mobilis. Mol. Gen. Genet. 209:374–381.
210. Wolff, J. B., and N. O. Kaplan. 1956. Hexitol metabolism in Escherichia coli. J. Bacteriol. 71:557–564.
211. Wong, P.-K., and E. L. Barrett. 1983. Aerobic and anaerobic alcohol dehydrogenases of Escherichia coli. FEMS Microbiol. Lett. 22:143–148.
212. Wu, L.-F., and M.-A. Mandrand. 1993. Microbial hydrogenases: primary structure, classification, signatures and phylogeny. FEMS Microbiol. Rev. 104:243–270.
213. Wu, L.-F., and M.-A. Mandrand-Berthelot. 1986. Genetic and physiological characterisation of new Escherichia coli mutants impaired in hydrogenase activity. Biochimie 68:167–179.
214. Wu, L.-F., M.-A. Mandrand-Berthelot, R. Waugh, C. J. Edmonds, S. E. Holt, and D. H. Boxer. 1989. Nickel deficiency gives rise to the defective hydrogenase phenotype of hydC and fnr mutants in Escherichia coli. Mol. Microbiol. 3:1709–1718.
215. Yerkes, J. H., L. P. Casson, A. K. Honkanen, and G. C. Walker. 1984. Anaerobiosis induces expression of ant, a new Escherichia coli locus with a role in anaerobic electron transport. J. Bacteriol. 158:180–186.
216. Zinoni, F., A. Birkmann, W. Leinfelder, and A. Bck. 1987. Cotranslational insertion of selenocysteine into formate dehydrogenase from Escherichia coli directed by a UGA codon. Proc. Natl. Acad. Sci. USA 84:3156–3160.
217. Zinoni, F., A. Birkmann, T. C. Stadtman, and A. Bck. 1986. Nucleotide sequence and expression of the selenocysteine-containing polypeptide of formate dehydrogenase (formate-hydrogen-lyase-linked) from Escherichia coli. Proc. Natl. Acad. Sci. USA 83:4650–4654.
218. Zinoni, F., J. Heider, and A. Bck. 1990. Features of the formate dehydrogenase mRNA necessary for decoding of the UGA codon as selenocysteine. Proc. Natl. Acad. Sci. USA 87:4660–4664.
Chapter18
