Fermentative Pyruvate and Acetyl-Coenzyme A Metabolism

TOC

doi: 10.1128/ecosal.3.5.3

R. Gary Sawers¹* and David P. Clark²

[SECTION EDITOR, AUGUST BÖCK]

Posted July 27, 2004

¹ Department of Molecular Microbiology, John Innes Centre, Norwich NR4 7UH, United Kingdom, and ² Department of Microbiology, Southern Illinois University, Carbondale, IL 62901

*Corresponding author: Phone: 44 1603 450750, Fax: 44 1603 450778, E-mail: This e-mail address is being protected from spambots. You need JavaScript enabled to view it

INTRODUCTION

Pasteur originally described fermentation 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 any organism that flourishes by this mode of growth. The enterobacteria, with their remarkably flexible metabolisms, are (together with clostridia), the classical organisms of bacterial fermentation. When grown on glucose in the absence of oxygen, Escherichia coli will excrete acetate, carbon dioxide, dihydrogen, ethanol, formate, lactate, and minor amounts of succinate. While succinate is derived from phosphoenolpyruvate (PEP), all other fermentation products originate from pyruvate.

Pyruvate can be reduced and excreted as D-lactate, or it can be cleaved homolytically to acetyl-coenzyme A (CoA) and formate (Fig. 1). Acetyl-CoA can then be converted to acetate through the successive action of phosphotransacetylase (PTA) and acetate kinase (ACK) to produce acetate and one molecule of ATP, as outlined in the following reaction sequence:

Fig. 1Fermentation pathways of E.coli, starting with glucose, glucitol, gluconate, or glucuronate, as examples for fermentable substrates. Depending on the substrate, either the Embden-Meyerhoff or the Entner-Doudoroff glycolytic pathway is used. Fermentation products are colored green; ATP and NAD ⁺are colored red. Key enzymes of fermentation are shown.

Pyruvate + CoA ↔ acetyl-CoA + formate (Δ G °' = −3.9 kcal/mol) (1)

Acetyl-CoA + P_i ↔ acetyl phosphate + CoA (Δ G °' = +2.2 kcal/mol) (2)

Acetyl phosphate + ADP ↔ acetate + ATP (Δ G °' = −3.1 kcal/mol) (3)

This succession of reactions is the backbone of the cellular machinery for the anaerobic life of the enterobacterial cell and represents the epitome of substrate level phosphorylation when the cells are growing on pyruvate, since there is no formal redox chemistry involved in the three reactions. In pyruvate metabolism, the E. coli cell is sustained anaerobically solely by the single ATP generated by reactions 1 through 3 (171). In bioenergetic terms, pyruvate is classified as a high-energy compound with an acetyl group transfer potential (Δ G °') (pyruvate hydrolysis to acetate and formate; calculated from the values given above and −31.8 kJ mol⁻¹ for ATP hydrolysis) of 51.9 kJ mol⁻¹ (224).

In order to ensure that reoxidation of NADH is maintained during fermentative growth on glucose as a carbon source, enterobacteria have evolved a second route of acetyl-CoA metabolism, which involves its successive reduction via acetaldehyde to ethanol (Fig. 1). The enzyme that catalyzes these reactions is alcohol dehydrogenase (AdhE), and it is also intimately involved in controlling the activity of the pyruvate-cleaving enzyme pyruvate formate-lyase (PFL). Consequently, depending on the oxidation state of the carbon substrate, the ratio of acetate to ethanol production varies.

This module will focus principally on the fermentation of E. coli and Salmonella, and in particular on the physiology, enzymology, and genetics of acetate, lactate, and ethanol production. A separate module will deal with formate metabolism. Since studies have revealed that E. coli and Salmonella are essentially indistinguishable with regard to their fermentative metabolisms, the module has been written with this in mind. Other genera of the enterobacteria, however, have evolved a specialized fermentative pathway not present in either E. coli or Salmonella enterica serovar Typhimurium that also initiates from acetyl-CoA. Thus, Enterobacter and Klebsiella produce the alcohol 2,3-butanediol, and we have included a section that discusses the 2,3-butanediol pathway in the light of energy generation and the maintenance of redox balance.

CONCEPT OF FERMENTATION BALANCE

Reducing Equivalents Produced

The oxidation of hexoses, such as glucose, to pyruvate by the Embden-Meyerhof-Parnas pathway generates two molecules of NADH (Fig. 1). In order to maintain glycolytic flux, the NADH must be reoxidized to NAD⁺, and during fermentation, this is achieved by depositing the reducing equivalents on metabolic intermediates, yielding reduced fermentation end products that are then excreted from the cell. The fermentation products of E. coli comprise a mixture of ethanol and acetic, formic (cleaved to H₂ + CO₂, which are derived from formate), lactic, and succinic acids (35, 207). These products have different oxidation states (Table 1), and by adjusting the ratios of the end products, E. coli modulates its metabolism to grow fermentatively on a variety of compounds, such as hexitols and hexonic and hexuronic acids, in addition to hexoses themselves (see Chapters Hexose/Pentose and Hexitol/Pentitol Metabolism and Catabolism of Hexuronides, Hexuronates, Aldonates, and Aldarates).

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 to 1,3-bisphosphoglycerate by glyceraldehyde-3-phosphate dehydrogenase (Fig. 1). More reduced sugar alcohols, e.g., glucitol (sorbitol), enter the cell via the PEP-sugar phosphotransferase system as sorbitol-6-phosphate, which is then oxidized before entering the Embden-Meyerhof-Parnas pathway as fructose-6-phosphate (see Chapter Glycolysis and Flux Control). The dehydrogenation of sorbitol-6-phosphate generates a further molecule of NADH, in addition to those produced during glycolysis (160, 242). Consequently, more reduced fermentation products need to be excreted during fermentation of glucitol 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 during glycolysis (1, 35, 207). Fermentation of gluconate or glucuronate generates only one molecule of NADH; however, the metabolism of glucuronate requires that it be reduced via a fructuronate intermediate to mannonate by D-mannonate dehydrogenase (81), thus oxidizing a molecule of NADH, which is generated during glycolytic degradation of the mannonate to two pyruvates. Therefore, conversion of glucuronate to two molecules of pyruvate is redox balanced (Fig. 1). However, fermentation of the last two compounds affords redox poise maintenance by reducing less metabolic intermediates and also conserves less energy during glycolysis (see below), since they are both metabolized via the Entner-Doudoroff pathway (Fig. 1). These examples show quite clearly that fermentation is a compromise: attempting to attain maximal energy conservation at the cost of upholding redox balance.

Reducing Equivalents Consumed

To attain balanced redox poise, all of the reducing equivalents generated during glycolysis must be accounted for by the products formed. Only a minor part of the reducing equivalents can be consumed anabolically. Instead, E. coli has several possibilities available for fermentative NADH reoxidation, and Fig. 1 illustrates this metabolic potential. From every molecule of pyruvate cleaved by PFL, one molecule of formate and one molecule of acetyl-CoA are formed. Therefore, maximally one-third of the carbon of glucose can be converted to formate. Formate is either excreted from the cell or decomposed to carbon dioxide and dihydrogen by the formate hydrogenlyase (FHL) complex (see Chapter Anaerobic Formate and Hydrogen Metabolism).

As mentioned in the introduction, acetyl-CoA has two alternative metabolic fates (Fig. 1). First, the energy of the thioester bond can be conserved in the form of ATP by the action of the PTA/ACK pathway, which yields acetate as a fermentation end product. However, the reducing equivalents formed during glycolysis are not consumed in this pathway. Second, the energy of the thioester bond of acetyl-CoA can be sacrificed by reducing it to acetaldehyde and further to ethanol through two successive, reversible dehydrogenation reactions catalyzed by AdhE (Fig. 1). Acetaldehyde is probably never released as an intermediate in vivo (but see reference 48) due to its potential toxicity. Therefore, using this pathway commits the cell to consuming four reducing equivalents (2 NADH) per acetyl-CoA, rendering ethanol the most highly reduced major fermentation product of E. coli.

The other possibility for disposing of reducing equivalents is formation of lactate or succinate (Fig. 1). The D-lactate dehydrogenase (D-LDH) reaction results in the reoxidation of one NADH, but like the AdhE reaction, it has the disadvantage that it squanders an energy-rich pyruvate molecule. Succinate normally comprises only 5 to 10 mol% (Table 2) of the fermentation products, and the vast majority produced by fermenting cultures is derived from PEP (35, 207). The first step in this route is the carboxylation of PEP, with a large proportion of the CO₂ arising from the FHL reaction. Hence, the availability of metabolic CO₂ limits the amount of succinate production, at least in laboratory cultures without added bicarbonate. 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; however, this is an unlikely alternative, first because it produces two reducing equivalents, and second, because the glyoxylate cycle is strongly glucose repressed (138) (see Chapter Two-Carbon Compounds and Fatty Acids as Carbon Sources).

Table 2Calculation of 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 D-LDH reaction is superfluous. Indeed, Clark and colleagues have demonstrated that mutants unable to produce the fermentative NADH-dependent D-LDH have no obvious anaerobic growth defects (145) (see below). This finding is also in accord with the regulation of D-LDH enzyme synthesis and activity, both of which are low during the initial phase of fermentation but are activated by a shift to low pH values. Hence, during the growth of E. coli in batch culture, lactate is the organic acid that appears last (215). The reasons that the D-LDH reaction has been conserved during evolution are probably the enhanced metabolic flexibility the reaction gives to the organism and, perhaps more important, the energy value of lactate (see Energy Balance below). Interestingly, E. coli B strains excrete large amounts of D-lactate when growing fermentatively with glucose (121), and E. coli K-12 strains can be engineered to conduct homolactate fermentation (254). This suggests that selective pressure may maintain the D-LDH reaction.

Studies with ack, adhE, and pta mutants have demonstrated the essential requirement for the derived enzymes to permit growth of E. coli on certain fermentable sugars. Mutants unable to synthesize AdhE do not grow anaerobically on glucitol, glucose, or gluconate but do ferment the highly oxidized glucuronate (43, 73). This reflects the fact that metabolism of glucuronate to pyruvate is redox balanced (Fig. 1). Neither ack nor pta mutants grow on any of the above-mentioned substrates (38, 73). However, strains defective in both pta and adhE or adhE and pfl regain the ability to ferment glucose (73, 171), and essentially, such double mutants perform lactate fermentation. It would be anticipated that pta or ack single 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 and suggests that there may be some common allosteric regulatory mechanism controlling the activities of the PFL, PTA, ACK, and AdhE enzymes. An important role of AdhE in regulating PFL activity (as PFL deactivase) has been demonstrated (102, 104).

Calculation of Fermentation Balance

Determination of the fermentation balance provides information on the catabolism of a particular sugar by a particular organism. It has been determined that only ~10% of the carbon source is assimilated due to the poor energy yields of fermentation. Therefore, fermentation balance studies are particularly useful for comparing and contrasting the metabolic fates of different sugars, or their derivatives, in different bacterial strains (37, 207).

The organism to be investigated is placed within a sealed vessel under an inert atmosphere, and fermentation is 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 (2). In vivo nuclear magnetic resonance spectroscopy has been developed as a useful method for examining the progress of fermentation (1, 145, 164, 165).

An example of a fermentation balance of E. coli grown with glucose is presented in Table 2. The sum of the oxidized and reduced products (redox balance) should equal the oxidation state of the substrate. In the example presented, the sum is +0.28, suggesting that not all products were efficiently recovered (Table 2). The carbon recovery is obtained by multiplying the number of 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 (plus CO₂) ratio was ~1:1:2:2 (Table 2). Hence, one-third of the carbon from glucose was metabolized to lactate. The total number of moles of ethanol plus acetate equals the total number of moles of formate plus CO₂ produced, which is in agreement with the anticipated stoichiometry from the cleavage of pyruvate by PFL (Fig. 1).

Based on 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 (1). 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. 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 is channeled through the PTA/ACK pathway to synthesize ATP (Fig. 1).

Energy Balance

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. Yet fermentation yields only two ATP per glucose from glycolysis (Fig. 1). Acetyl-CoA is therefore used as an additional "energy-rich" intermediate during fermentation (224), potentially yielding 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 regenerates only one additional ATP per glucose, resulting in a total of three ATP per glucose. The fermentation of more reduced substrates necessitates a higher proportion of ethanol production with concomitantly less ATP regenerated via acetate kinase (Fig. 1). More oxidized substrates, on the other hand, such as gluconate or glucuronate, should permit more ATP to be generated by substrate level phosphorylation, but because these substrates are metabolized by the Entner-Doudoroff glycolytic pathway, one less ATP is regenerated during their degradation to pyruvate.

There is also a considerable energy potential stored in the reduced organic acids produced. Michels 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 (156). 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. Experimental validation of this proposal has been presented for L-lactate efflux employing E. coli membrane vesicles (223).

PYRUVATE CLEAVAGE

LDH

Lactic acid is produced by reducing pyruvate and occurs in a wide range of organisms when they grow under fermentative conditions. Some organisms produce L-lactate, whereas others produce D-lactate. Mammalian muscles, like vertebrates and plants in general, produce L-lactic acid when in oxygen debt. Generally, vertebrates and plants produce L-lactate when in oxygen debt, and prokaryotes usually produce either L- or D-lactate, with a few organisms able to produce both. For example, Lactobacillus plantarum cells contain distinct L- and D-LDHs in the same cell (218).

Biochemistry of LDH.

E. coli contains three enzymes that interconvert pyruvate and lactate. Two of these are membrane-bound, flavin-linked pyruvate oxidoreductases that convert lactate to pyruvate during respiratory metabolism in the presence of oxygen or other electron acceptors, such as nitrate. These enzymes are specific for either the L isomer or the D isomer and allow the use of lactate as a carbon and energy source. The third enzyme is a soluble, NAD⁺-linked LDH that converts pyruvate to the D-lactate during fermentation.

E. coli does not contain a soluble NAD⁺-linked L-LDH. An early report of L(+)-lactate being generated by the fermentative LDH of E. coli and secreted into the medium (220) has been corrected, showing specificity of the enzyme for D-(–)-lactate (221). The fermentative D-LDH as purified from E. coli strain B consists of four identical subunits and exhibits a molecular mass of 115 kDa. It is inhibited by thiol reagents, which react with an essential cysteinyl residue (200).

The D-LDH is allosterically activated by its substrate, pyruvate. Consequently, there are two binding sites for pyruvate, representing the active site and an allosteric binding site. The difference between the two sites is illustrated by their varied specificities for different pyruvate analogues. Thus, oxamate acts as a competitive inhibitor for pyruvate reduction but has no effect on allosteric activation. Conversely, α-ketobutyrate does not inhibit the enzyme reaction but does mimic pyruvate in activating the enzyme (222). ATP in millimolar concentrations inhibits D-LDH, probably by competing with NAD⁺.

The affinity of the enzyme for pyruvate is low—the active site has a K_m value of ~7 mM, and the enzyme is only fully activated allosterically in the presence of ~5 mM pyruvate. The K_m value for pyruvate increases at higher pH, with concomitant decrease of V _max; hence, the activity of D-LDH is higher at lower pH values, and more lactate is actually produced under such conditions (222). The high K_m value of LDH means that it is poorly equipped for competition with PFL for pyruvate (the K_m for pyruvate is 2 mM). Consequently, even when LDH is 10-fold overproduced, lactate synthesis only doubles (251). Adding high levels of exogenous pyruvate to fermenting cultures leads to an increase in the intracellular pyruvate pool and increased lactate formation (250). This is presumably due to allosteric activation of LDH as a result of increased cytosolic pyruvate levels.

The allosteric regulation of the E. coli D-LDH is quite distinct from that of the LDHs typically found in L-lactate-producing lactic acid bacteria. The L-LDH of these bacteria is activated by fructose 1,6-biphosphate rather than by pyruvate (150).

Genetics and Sequence Relationships of LDH.

A defect in the D-(−)-LDH alone has no known growth phenotype, since fermentation may continue via production of ethanol-acetate. However, mutants impaired in D-LDH activity were isolated by starting with a strain that lacked functional PFL and subsequently screening for mutants no longer capable of growing fermentatively (145). These ldhA mutants had very low or no detectable NAD⁺-linked LDH activity and no longer produced lactate as a fermentation product (145).

The ldhA gene was mapped at 31.0 min on the E. coli chromosome and cloned and sequenced (26, 145). The fermentative D-LDH of E. coli encoded by this gene consists of 329 amino acids and has a molecular mass of 36,532 Da. It is highly similar to orthologous enzymes from other enteric bacteria, including Salmonella, Shigella, and Yersinia, and is less closely related to D-LDHs from a wide range of other bacteria and even some lower eukaryotes, such as Mastigamoeba and Schizosaccharomyces. Strong overproduction of plasmid-borne D-(−)-LDH caused growth defects, probably due to conversion of the cytosolic pyruvate pool to lactate (26). Supplementation with pyruvate or some other three-carbon compounds partially relieved these growth defects.

Complementation of the anaerobic growth defects of ldhA pfl double mutants of E. coli has been used successfully to clone LDH genes from a variety of other organisms. Genes for several D-(−)-LDHs from Lactobacillus species (12, 13), as well as a gene for fructose 1,6-biphosphate-activated L-(+)-LDH from Clostridium sp. (40), have been cloned and shown to function efficiently in E. coli.

The D-LDHs form a subgroup within a larger family of D-specific 2-hydroxyacid dehydrogenases. This family includes both degradative and biosynthetic enzymes. Examples are D-hydroxy-isocaproate dehydrogenase, D-glycerate dehydrogenase, D-3-phosphoglycerate dehydrogenase (the serA gene product in E. coli), and D-erythrose-4-phosphate dehydrogenase (the pdxB gene product in E. coli).

E. coli contains two other less characterized D-specific 2-hydroxyacid dehydrogenases that show some sequence similarity to the ldhA gene product. These are the products of the ycdW and yiaE genes. Both derived enzymes act on glyoxylate and hydroxypyruvate. Although YcdW preferentially uses glyoxylate, hydroxypyruvate is the preferred substrate of YiaE (162).

Although all known L-LDHs from any organism are highly similar, the sequences of the D-LDHs are completely unrelated to these enzymes, except for a G-X-G-X-X-G motif close to the N terminus in L-LDHs (19, 133, 218) but around position 150 to 160 in D-LDHs (19, 218). This motif is common to most NAD-linked dehydrogenases (19), as part of the β α β fold, which constitutes the NAD⁺-binding domain (57, 240).

Possible Energy Conservation from Lactate Export.

The branched fermentation pathway that generates acetate plus ethanol is generally regarded as superior to the lactate route from the viewpoint of energy generation, since one ATP is regenerated at the ACK step. It has been suggested, however, that some energy might be produced by efflux of lactate. Any metabolite that is present in high concentration inside the cell and in low concentration outside could in theory release energy as it moves outward. The generation of proton motive force requires coupling solute efflux to proton movements via a suitable permease.

Membrane vesicles of E. coli have been shown to generate proton motive force when loaded with solutes, such as lactose, proline, or lactate. For L-lactate, the H⁺/L-lactate stoichiometry of symport varies between 1 and 2, depending on the external pH. This change in stoichiometry is most likely caused by a protonation-deprotonation of the lactate carrier protein in response to the external pH. Unfortunately, these experiments were done in E. coli strain ML 308-255, which is not a K-12 derivative, and the L isomer of lactate was used (223). The question of whether the efflux of fermentatively generated D-lactate provides E. coli with extra energy remains open. In homolactate-fermenting Streptococcus cremoris, the efflux of L-lactate has indeed been shown to contribute to proton motive force generation (114).

Uptake of L-lactate, D-lactate, and glycolate during aerobic growth on these substrates is mediated by the LldP or GlcA permease (161). These enzymes are part of the metabolic systems for L-lactate and glycolate, respectively. No separate carrier is presently known for D-lactate. However, an uncharacterized gene (referred to as hslJ) (34, 131), immediately downstream of ldhA and possibly in the same operon, codes for a hydrophobic protein that may be a candidate for a lactate export carrier.

Regulation of LDH.

The fermentative D-LDH is regulated at several levels. The enzyme itself is regulated allosterically in response to pyruvate levels, as discussed above (222). In addition, the expression of the ldhA gene is induced by acidification of the medium during anaerobic growth (93, 145). Regulation occurs at both the DNA and RNA levels, although the details are only partly understood at present. Regulation of the D-LDH has turned out to be surprisingly complex, and the ldhA gene itself displays several transcriptional start sites that have yet to be fully characterized. It should be noted that the LdhA protein was characterized in E. coli B, whereas the regulatory studies were performed in E. coli K-12. It is not known whether there are any significant differences in the behavior of LdhA between these strains of E. coli.

During nitrate respiration, accumulation of lactate is greatly decreased (11). D-LDH is inhibited in its activity by nitrate and nitrite (247), but not in its synthesis, as indicated by unchanged protein content. Competition for NADH oxidation by anaerobic respiration with nitrate is probably another factor limiting lactate production under these conditions. In accord with these observations, it was found that narL or fnr mutants are not affected in expression of ldhA.

In E. coli, lactate is a subsidiary fermentation product that is produced late during fermentation after substantial acidification has already taken place due to the generation of a mixture of acetic acid, formic acid, and ethanol. Not surprisingly, transcription of the ldhA gene of E. coli is induced ~10-fold by a combination of anaerobiosis and acidity but not significantly by either condition alone (35, 145). Expression of ldhA is not induced by membrane-permeant weak acids, which implies that expression is not directly controlled by the internal pH. Moreover, only weak acidic pH induction takes place during anaerobic growth on glycerol plus fumarate (93). Indeed, the addition of pyruvate to the medium caused a significant increase in expression of the ldhA gene (93).

The arcAB two-component regulatory system has a major effect on ldhA expression at the transcriptional level. Mutations in either arcA or arcB result in a massive drop in ldhA expression, implying that the ArcA protein behaves as an activator of ldhA (93). In accord with this, there is an ArcA-binding site at positions −64 to −73 relative to the transcription initiation site of the ldhA gene (M. R. McGehee and D. P. Clark, Abstr. 102nd Gen. Meet. Am. Soc. Microbiol., abstr. K-152, p. 293, 2002). It has been shown that synthesis of the reduced fermentation products ethanol and succinate is not significantly induced until oxygen concentrations drop below the microaerobic level, i.e., below a partial pressure of ~100 Pa (11). However, lactate is already produced in significant amounts under microaerobic conditions, which corresponds well with the known role of ArcA in regulating gene expression under these conditions. These findings, however, are in conflict with the observations of Riondet et al. (183). These authors compared anaerobic cultures grown at different redox potentials. They found that the ratio of ethanol to lactate was high at a culture redox potential of −100 mV, while LDH activity increased three to sixfold under more reducing conditions (E₀ = −320 mV).

The cyclic AMP (cAMP)-cAMP receptor protein system also affects ldhA expression, although in a somewhat puzzling way. Anaerobically, LdhA levels are higher in cells grown on glucose than in those grown on other sugars, presumably due to more rapid acidification and/or pyruvate accumulation. However, in cya or crp mutants, ldhA expression is reduced ~5-fold (McGehee and Clark, Abstr. 102nd Gen. Meet. Am. Soc. Microbiol.), which is rescued by the addition of pyruvate to the cultures. Sequence analysis reveals a Crp binding site at positions −10 to −30 upstream of the transcription initiation site of the ldhA gene. Such a location is highly unusual, and whether this site is functional is as yet uncertain.

Translational ldhA-lacZ fusions consistently showed ~10-fold-higher expression than transcriptional fusions, implying some posttranscriptional effects on ldhA regulation (93). This effect was localized to a 60-bp region at the 5' end of the ldhA gene. Removal of this segment reduced expression by ~10-fold (92). A probable candidate mediating this effect is the small regulatory RNA ryhB(synonym, sraI), which affects expression of several genes involved in Fe transport and metabolism (143), as well as ldhA (S. Gottesman personal communication). Although ryhB has a negative effect on most of its target genes, the reverse is true for ldhA. Knockouts of ryhB result in decreased expression of ldhA-lacZ translational fusions (but have no effect on transcriptional fusions), implying that ryhB enhances ldhA-lacZ translation (M. R. McGehee and D. P. Clark, unpublished data). Expression of ryhB itself is controlled by the Fur repressor in response to iron availability (143). The cellular ryhB level is adjusted by degradation via RNase E and protection from degradation by the RNA-binding protein Hfq (142, 158).

Why ldhA should be regulated by the Fur/ryhB system is uncertain. However, one possible reason is that D-LDH does not contain Fe, whereas AdhE, one of the major proteins in fermenting cells, is an Fe-containing protein, as are some of the enzymes involved in succinate formation. Thus, one possibility is that when faced with a Fe shortage, AdhE synthesis is reduced and lactate fermentation is used instead. Much further investigation is needed in this fascinating area.

Role of D-LDH in Metabolism.

D-LDH plays two main metabolic roles. Under anaerobic conditions, it provides an alternative fermentation route for NADH reoxidation. Late in fermentation and/or when the medium has become acidified, E. coli switches from ethanol-acetate production to lactate fermentation. Admittedly, this results in decreased acid production, but additional factors may be involved. Only two lactate molecules are generated per glucose fermented instead of two molecules of formate plus one molecule of acetate. However, in real life, E. coli is a minor member of the intestinal population and would have little overall effect on the pH of its surroundings. Furthermore, formic acid can be eliminated by conversion to carbon dioxide plus hydrogen, whereas lactate remains in the medium. Most of the information available applies to the K-12 strain of E. coli, which is tacitly assumed to be typical in its fermentative behavior. However, although little detailed information is available, it is clear that many wild strains of E. coli differ markedly from the K-12 pattern. For example, E. coli strain B tends to ferment glucose to high levels of lactate and makes almost no ethanol-acetate early in fermentation, even at neutral pH (121). Nonetheless, E. coli B does make high levels of ethanol-acetate when grown on xylose, showing that the ethanol-acetate pathway is intact (121) but apparently is used during growth on less favored sugars.

Although induced anaerobically, the basal level of D-LDH present under aerobic conditions is still significant. Given the allosteric properties of the enzyme, D-LDH is active only when pyruvate levels are relatively high. Thus, it seems likely that D-LDH is involved in removing pyruvate if the concentration exceeds a threshold level as a result of some metabolic disturbance or block (69, 254). For example, it appears that quinone-deficient mutants of E. coli that are unable to respire rely on D-LDH to metabolize pyruvate aerobically (11).

PFL

The oxidative decarboxylation of pyruvate by the pyruvate dehydrogenase complex (PDHC) generates one NADH per molecule of pyruvate. Consequently, it was originally concluded that PDHC is functionally restricted to respiratory metabolism (72, 97). More recent studies, however, in which the metabolic flux has been determined through PDHC as a function of cellular redox potential (the NADH/NAD⁺ ratio), indicate that PDHC activity, and to a large degree its synthesis, is controlled by the redox state, as manifested by the prevalent concentrations of the NADH-NAD⁺ redox couple (2, 49). The activity of the PDHC complex is negatively regulated by NADH (76); therefore, as the NADH/NAD⁺ ratio increases when the cells become more anoxic, a transition occurs in which PDHC is inactivated and the anaerobic alternative enzyme PFL becomes activated (2, 49). When the NADH/NAD⁺ ratio is high—for example, during fermentation—nonoxidative cleavage of pyruvate to acetyl CoA and formate by PFL is of clear benefit to E. coli (103, 111, 190, 198).

PFL enzyme activity is strictly controlled by a remarkably sophisticated interconversion cycle (103, 111, 113). PFL must be activated by the introduction of a stable glycyl radical into the polypeptide chain (Fig. 2). This reaction is catalyzed by an iron-sulfur cluster-containing activating enzyme, which requires reduced flavodoxin and S-adenosylmethionine (AdoMet) as substrates. The active glycyl radical-containing form of PFL is extremely sensitive to dioxygen, and consequently, removal of the radical and conversion of the enzyme to the inactive, dioxygen-stable species is catalyzed in a controlled manner by another enzyme, which is identical with the fermentative acetyl-CoA reductase-AdhE.

Fig. 2PFL posttranslational interconversion cycle. The PFL-activating (PFL-AE) and –deactivating (AdhE) enzymes are shown in red. The radical-free form of the enzyme is designated PFL-H, and the radical-containing form of PFL is shown as PFL ^•. Fd, ferredoxin; Fld, flavodoxin; FldH ₂, dehydroflavodoxin.

The redox status of the cell clearly is a key function in controlling which of the two enzymes, PDHC 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 PDHC complex are auxotrophic for acetate when grown aerobically but are prototrophs during fermentative growth (120). On the other hand, pfl mutants are prototrophic during aerobic growth but grow very poorly by fermentation if the culture medium is not supplemented with acetate (49, 97, 145, 228). Interestingly, when cultures use alternative electron acceptors, such as nitrate (+430 mV for nitrate-nitrite), with redox potentials lying between the extremes found during aerobic (+820 mV for water-oxygen) and fermentative (−420 mV for hydrogen-proton) growth, then either of the two enzymes can be functional in metabolizing pyruvate (97). Moreover, recent steady-state culture analyses have demonstrated that in the low microaerobic range of dioxygen concentration, the two enzymes have overlapping activities (2, 49). This indicates that, despite the presence of low levels of dioxygen in the environment, the PFL enzyme is in its active, radical form. The resolution of this apparent paradox is thought to be due to the scavenging of intracellular dioxygen by respiratory cytochrome d oxidase, the synthesis of which, like that of PFL, is induced microaerobically (2). The role of cytochrome d oxidase in preventing the deleterious effects of dioxygen inactivating the radical-bearing PFL species is analogous to that proposed for the same enzyme in respiratory protection of the dioxygen-sensitive nitrogenase enzyme in the obligate aerobe Azotobacter vinelandii (175).

Structure of PFL.

PFL is a homodimer comprising two 759-amino-acid polypeptide chains (103). The key feature of PFL is that it uses radical-based chemistry to perform reversible pyruvate cleavage (110, 232). A series of detailed biochemical studies led to the proposal of a catalytic mechanism for the enzyme, but it has been ultimately the elucidation of the structure of the PFL enzyme that has provided most insight into the catalytic mechanism (Fig. 3).

Fig. 3(A) Structure of PFL monomer with pyruvate and coenzyme A bound. The locations of pyruvate and CoASH are shown by arrows. Residues Cys418, Cys419, and Gly734 are shown as ball-and-stick representations, with sulfurs shown in yellow. The Protein Data Bank accession code is 1h16. (B) Juxtaposition of key active-site residues in PFL, Gly734, Cys418, and Cys419. The sulfurs of the cysteinyl residues are shown in yellow. The arginines hydrogen bond with the pyruvate substrate. The Protein Data Bank accession code is 1h16.

The structure of PFL revealed that the enzyme is related to ribonucleotide reductases (RNR), in particular, to the class III anaerobic isoenzyme (9, 126, 134). Class III RNR employs a glycyl radical-based mechanism to reduce ribonucleotides to deoxyribonucleotides, required for DNA synthesis with formate as the actual reductant (212). In both PFL and RNR, the glycyl radical is formed by removal of a hydrogen atom from glycine, with the radical being delocalized on the neighboring carboxamides of the polypeptide backbone. The structure of PFL includes a 10-stranded α/β barrel structure comprising an antiparallel arrangement of two parallel 5-stranded β-sheets (9). Inside this barrel structure, there is a protruding hairpin, which has the adjacent centrally located Cys418 and Cys419 residues at its tip. Approaching this loop from the C-terminal end of the polypeptide and the other side of the barrel is a second hairpin loop structure with the Gly734 residue located at its tip. The juxtaposition of the two loops joins Gly734 and Cys419 at a distance within 3.2 Å (Cα distance, 4.8 Å) of each other and thus optimally positions the amino acids for hydrogen atom transfer from the thiol group of Cys419 to the glycyl radical initiating the reaction (Fig. 3). In addition, the adjacent Cys418 is also in close enough proximity to Cys419 to participate in radical transfer reactions and completes the catalytic triad.

The active site of PFL is hydrophobic and very compact, which presumably is conducive to the persistence of the radical (103). The compact structure, however, raises interesting questions regarding how the PFL-activating enzyme (PFL-AE) abstracts the hydrogen from Gly734 and how the Gly734 radical is quenched by the multimeric AdhE protein (Fig. 2).

In contrast to PFL, the active site of class III RNR has a more open structure (56, 134, 212). Intriguingly, the Cα atoms of 451-amino-acid residues in PFL superimpose (root mean square deviation, 3.1 Å) on equivalent (not necessarily identical) residues in NrdD (a class III RNR), including those present at the base of the barrel structure (113). It is notable that only weak similarity of PFL and NrdD is detectable at the primary structural level. These findings suggest that PFL and RNR evolved divergently from a common ancestor, but a long, parallel evolutionary history has led to preservation of a common overall structure.

Substrate Binding.

The original PFL structure solved by Becker et al. (9) had the pyruvate analogue oxamate in both active sites of the dimer. More recent cocrystal structures of the enzyme have pyruvate in the active sites and reveal that oxamate and pyruvate adopt similar conformations (10, 128). Interestingly, there are no major structural changes upon substrate binding. Perhaps the most significant aspect of pyruvate binding is the fact that the carbonyl carbon of pyruvate is located 2.6 Å from the Sγ of Cys418, and this has important implications for the catalytic mechanism (see below). The carboxylate oxygens form a salt bridge with the guanidino group of Arg435, while the carbonyl oxygen is hydrogen bonded to Arg176 and a water molecule (Fig. 3B). Finally, the substrate has a planar orientation and is sandwiched between the aromatic rings of Phe432 and Trp333. The Sγ of Cys418 is inserted between the pyruvate plane and Trp333, making Cys418 an obvious candidate for the location of a thiyl radical, attacking the carbonyl carbon of pyruvate (9).

The CoA-Binding Site.

The PFL reaction comprises two half-reactions, the second of which involves an S-acetyl transfer. Subsequent to C-C bond cleavage, the acetyl group is covalently attached to Cys418, and transfer to CoA has been proposed to occur through a radical-based mechanism. This is also consistent with the fact that the active site is hydrophobic and is unlikely to support a heterolytic thiol transfer reaction. Consequently, the location of the CoA-binding site is important with regard to this proposed mechanism. Becker and Kabsch (10) determined the structure of a PFL cocrystal with pyruvate and CoA and found a binding site for CoA in each subunit. Notably, CoA binding did not significantly perturb the conformation of the pyruvate-binding site. The site is located near the subunit interface, and surprisingly, the thiol of CoA is at a distance of 30 Å and the adenine is ~15 Å from the active site (Fig. 3A). However, the thiol of CoA present in the crystal is in the syn conformation, and the authors propose, based on model calculations, that upon completion of the first half-reaction, a conformational change must occur in the protein, which would result in CoA adopting an anti configuration. The postulated rotation of the bound CoA around the N-glycosidic bond, while keeping the adenine in place, would be compatible with the ribose-pantetheine moiety stretching like a finger into the active site, placing the thiol group of CoA in a suitable location to accept the acetyl group from Cys418.

Catalytic Mechanism.

The historical aspects of the elucidation of the PFL reaction mechanism are detailed in several reviews (103, 113, 213). We will attempt to focus on the salient features of the mechanism as proposed, based on recent structural information.

Essentially, PFL follows a "ping-pong" reaction mechanism, and the two half-reactions are shown in Fig. 4. The catalytic efficiency of the overall reaction is 770 s⁻¹ for the forward reaction and 260 s⁻¹ for the reverse reaction, values which compare very favorably with β-keto thiolase-catalyzed cleavage of C-C bonds following an ionic reaction mechanism (107). After the early identification of the involvement of radical chemistry (110) and identification of Gly734 as the site of radical storage (232), the interpretation of inhibitor studies and isotope exchange analyses led to proposals for the catalytic mechanism. It was the more recent structural studies, however, in particular those with substrates or substrate analogues bound, that yielded a detailed proposal of the likely mechanism of PFL (9, 10, 126, 128). Both Cys418 and Cys419 are required for activity, and the proximity of Cys418 to the pyruvate carbonyl in the PFL structure immediately suggested that the acetyl of the acetyl-enzyme intermediate identified 30 years ago (107) is located on this residue. The proximity of Cys419 to both Gly734 and Cys418 means that it is likely to function as a radical relay station between the glycyl radical and a catalytic Cys418 thiyl radical (Fig. 3B and 4). The proximity of Cys419 to Gly734 was also predicted from H exchange studies (170). Based on genome sequence analyses and other studies (reviewed in reference 113), the databases reveal that, apart from PFLs from various sources and the α-ketobutyrate lyase, TdcE, from E. coli, all glycyl radical enzymes catalyzing other or unknown reactions have a single cysteinyl residue in their active sites. The thiolytic cleavage reactions catalyzed by PFL and TdcE are reversible, while the chemically completely unrelated reactions catalyzed by other glycyl radical enzymes, such as class III RNR and, for example, benzylsuccinate synthase from Thauera aromatica (129), are irreversible. The two adjacent cysteinyl residues of PFL permit hydrogen atom shuttling and may be used as a signature which indicates the PFL reaction type and reversibility. Therefore, unknown gene products should be annotated as putative PFLs only when these two cysteines are present.

Fig. 4Proposed catalytic mechanism of PFL. The three amino acids Gly734, Cys418, and Cys419 involved in radical transfer and catalysis are shown, and the red dots indicate the locations of the radicals. Upon its formation, formate leaves the active site and is replaced by acetyl-CoA.

In the first half-reaction of the proposed mechanism, the radical is passed from Gly734 via Cys419 to Cys418, where homolytic C-C bond cleavage of pyruvate occurs, yielding a bound acetyl moiety and a formyl radical (Fig. 4). The formyl radical then abstracts a hydrogen from Cys419 to regenerate a Cys419 thiyl radical. Due to the hydrophobic nature of the active site in PFL, it is unlikely that thioester transfer from Cys418 to CoA occurs by an ionic mechanism but rather by another homolytic process involving a CoA-based thiyl radical, as shown in Fig. 4. A theoretical study has determined that the energy barrier of a thiyl reacting with either pyruvate or an acetyl thioester is low (12 kcal/mol) (82), and Knappe et al. (109) have determined that although the acetyl transfer can occur with the nonradical form of PFL, it is ~10⁵ times faster with the radical form of the enzyme.

In the second half-reaction, therefore, it is proposed that Cys419 abstracts a hydrogen atom from CoA to yield a CoA radical intermediate (Fig. 4). The CoA radical then accepts the acetyl group from Cys418 and completes the reaction cycle. It is still open to speculation whether the glycyl radical is regenerated or whether Cys418^• directly initiates a new catalytic cycle. The latter scenario might be in accord with the high catalytic constants of the enzyme.

As mentioned above, the findings of inhibitor studies are entirely consistent with the proposed mechanism. For example, Plaga et al. (173) determined that the pyruvate analogue acetyl phosphinate and the formate analogue hypophosphite both result in the dead-end product 1-hydroxyethyl phosphonate thioester linked to Cys418. More recently, the pyruvate analogue methacrylate was shown to derivatize Cys418 with an S-(2-carboxy-(2S)-propyl) substitution, which results from a nucleophilic attack by a thiyl radical (174). Significantly, this dead-end enzyme retains the glycyl radical.

The radical in PFL is stable indefinitely anaerobically and at low temperature, and this remarkable stability is attributed to the nature of the PFL active site and to the fact that the radical is buried within the protein, ~20 Å from the protein surface. In the presence of dioxygen, however, the radical has an extremely short half-life of ~10 s at 0°C. Dioxygen causes cleavage of the polypeptide backbone at the N-Cα of Gly734, resulting in an ~3-kDa C-terminal peptide with an oxalyl residue at its N terminus (232). This study, along with corroborative mutagenesis studies, originally defined the location of the radical in PFL.

Detailed spectroscopic analyses of PFL inactivation by dioxygen have provided convincing evidence of protein-based sulfinyl and peroxyl radicals as intermediates in the inactivation process (179, 253). The studies provided evidence that dioxygen quenches the glycyl radical but also demonstrated that the short-lived peroxyl radical of Gly734 reacted with Cys419 and not Cys418 to form the longer-lived sulfinyl radical. Furthermore, using acetylated enzyme, these studies confirmed that the acetyl intermediate was located exclusively on Cys418.

Two important unresolved questions are not addressed by the present structural data. The first is that PFL exhibits half-of-the-sites activity. In other words, in the PFL dimer, only one of the Gly734 residues carries a radical in the resting active state of the enzyme. This observation is corroborated by dioxygenolytic cleavage of PFL, which results in 1:1 stoichiometry between full-length and cleaved polypeptides (232). In the crystal structures of nonactivated PFL, both sites are occupied by substrate. Clearly, this problem will be resolved by determining the structure of the radical form of PFL. The second question concerns the mechanism of enzyme-based radical introduction into, and removal from, PFL. Since Gly734 is buried well below the enzyme's surface, a significant conformational change must be invoked to account for both processes.

Posttranslational Modification of PFL.

Introduction of the radical into PFL is controlled by an exquisitely poised, redox-sensitive, posttranslational interconversion system (Fig. 2). The radical insertion reaction catalyzed is E-H + AdoMet + e⁻ → E^• + 5'-deoxyadenosine + methionine, where E-H is inactive PFL and E^• represents the active glycyl radical species.

When synthesized, PFL contains no glycyl radical and is enzymically inactive. The enzyme must undergo reductive activation, which involves direct hydrogen atom abstraction from Gly734 (103). The requirements for hydrogen atom abstraction include AdoMet, reduced flavodoxin I (FldI), and the iron-sulfur protein PFL-AE. AdoMet was the first cofactor to be identified as being involved in enzyme activation (108). Unlike most reactions involving AdoMet, no alkylation of the substrate occurs, but rather AdoMet is irreversibly cleaved during the activation reaction, yielding methionine and 5'-deoxyadenosine. It was demonstrated that an intermediary 5'-deoxyadenosyl radical abstracts the proS hydrogen directly from Gly734 (62).

As mentioned above, the radical form of PFL is extremely labile in the presence of dioxygen, so when the E. coli cell encounters oxygen, the radical is reduced and PFL is converted by AdhE to the oxygen-stable, inactive species. This reaction requires CoA, NAD⁺, and reductant (Fig. 2).

Generation of Reduced Flavodoxin.

E. coli has two flavodoxins and an adrenodoxin-like ferredoxin. Although ferredoxin was identified and copurified in the early study by Vetter and Knappe that determined FldI to be the electron donor required for PFL activation (229), ferredoxin does not substitute for FldI. However, a recent biochemical study showed that the semiquinone form of FldI obtains an electron from reduced ferredoxin (Fig. 2) (237). FldI has a flavin mononucleotide cofactor, and the redox couple of the dihydro and semiquinone forms is between −440 and −455 mV (229). Structural analysis of FldI from E. coli reveals that the flavin is tightly, but noncovalently, bound within the protein, and a hydrophobic surface near the cofactor has been proposed to be involved in selective protein-protein interaction (87).

The two known electron donor systems to FldI are NADPH-ferredoxin/flavodoxin oxidoreductase (NFOR) and pyruvate-ferredoxin/flavodoxin oxidoreductase (PFOR) (Fig. 2). NFOR is actually a comparatively poor electron donor to FldI (235), perhaps suggesting that in vivo PFOR is the principal FldI reductase for PFL activation. The structure of NFOR has been determined to 1.7-Å resolution (90). Little information is available concerning PFOR from E. coli.

PFL-AE.

PFL-AE is a 28-kDa monomeric iron-sulfur protein. The primary sequence of the protein has an N-terminal Cys-X₃-Cys-X₂-Cys motif, which is also found in the β protein subunit of class III RNR (Fig. 5) (219) and a growing family of other enzymes catalyzing unrelated AdoMet-dependent radical reactions, the "SAM radical generators" (206). The requirement for iron in the PFL activation reaction had been recognized for a considerable time (110, 112). However, it was only after anaerobic isolation that it became clear that the three N-terminal cysteinyl residues coordinate an unusual [4Fe-4S]²⁺ cluster (21, 22, 117). In the absence of the substrate AdoMet, it is unclear what acts as a ligand to the fourth Fe in the cluster. Incubation of PFL-AE with AdoMet results in alteration of the electron paramagnetic resonance spectrum of the protein from an axial to a rhombic signal (117), which clearly indicates that AdoMet coordinates the fourth unique Fe in the cluster. This has been confirmed by Mössbauer spectroscopy (63, 115).

Fig. 5[4Fe-4S] ⁺cluster of PFL-activating enzyme. (Left) Alignment of amino acid sequences around the three conserved cysteinyl residues proposed to coordinate the iron-sulfur cluster among the recognized gene products of glycyl radical-activating enzymes of E.coli. The numbers above the first amino acids indicate the locations in the polypeptide chain. NrdG T4 is the activating enzyme subunit of class III anaerobic ribonucleotide reductase from bacteriophage T4 (accession number P07075); NrdG is the activating enzyme subunit of class III anaerobic ribonucleotide reductase from E.coli (accession number NP_418658); PFL-AE is the PFL-activating enzyme from E.coli (accession number NP_415422); PFLE is the putative activating enzyme of the PFL-like enzyme PFLF (see the text for details) from E.coli (accession number P75794); PFLC is the putative activating enzyme of the PFL-like protein PFLD from E.coli (accession number P32675). (Right) Model of [4Fe-4S] cluster of PFL-AE and simplified mode of AdoMet binding and homolytic cleavage based on studies presented in reference 237.

Proposed Mechanism of 5-Deoxyadenosyl Radical Generation.

The new role for iron-sulfur proteins in the reductive cleavage of AdoMet has been reviewed recently (32, 59, 63).

Data obtained from a combination of selective labeling, together with electron paramagnetic resonance, electron nuclear double resonance, Mössbauer, and extended X-ray absorption fine-structure spectroscopies with bound Se-AdoMet have provided a plausible mechanism for PFL radical generation (42, 115, 117, 235, 236). In this mechanism, the unique Fe in the [4Fe-4S] cluster of PFL-AE has a novel role in substrate binding, and the electron is transferred to AdoMet via a sulfide of the cluster (Fig. 5).

Although AdoMet will interact with both the oxidized and reduced forms of the cluster (115), Külzer et al. (117) demonstrated that the [4Fe-4S]¹⁺ → [4Fe-4S]²⁺ oxidation accompanies AdoMet cleavage. Krebs et al. (115) determined that the amino, the carboxylate, and the ribose moieties of AdoMet were candidates for Fe ligation, which suggested that the Fe was not involved directly in the cleavage of AdoMet. Electron nuclear double resonance spectroscopy then determined that the methyl carbon of AdoMet was within 4 to 5 Å of one of the Cys-liganded Fe atoms, which indicates that the methionyl and not the adenosyl moiety is proximal to the unique coordinating Fe (235). Recently it has been determined that both the amino and the carboxylate groups of AdoMet are coordinated with the unique Fe in the cluster (42, 236). This novel anchoring role of the [4Fe-4S] cluster implies that substrate reduction occurs via a formal sulfide-sulfonium interaction (Fig. 5). This also would indicate that sulfur-based redox chemistry directs the irreversible C-S bond cleavage. This proposed reaction mechanism contrasts with the proposed Fe-based redox chemistry of AdoMet reductive cleavage in the activation of the analogous SAM radical generator lysine 2,3-amino mutase (59, 63).

It is important to emphasize that the nonradical form of PFL, or at least a peptide resembling the Gly734 site, is a prerequisite for AdoMet cleavage by activating the enzyme (62, 231). Moreover, pyruvate, or the oxamate analogue, is an essential allosteric effector for radical introduction. These observations have major implications for the delivery of the 5'-deoxyadenosyl radical to the active site of PFL. Only pyruvate-bound, inactive PFL triggers AdoMet cleavage and radical release. The structural data on PFL indicate that pyruvate binds only at the active site, and this indicates that the 5'-deoxyadenosyl radical is delivered directly to the active-site glycyl residue. Moreover, Becker and Kabsch have reported that 5-deoxyadenosine binds to PFL at the same site as the adenosyl moiety of CoA (10).

The protein-protein interactions involved in the radical introduction reaction are unclear. It can be envisaged that the chain of events includes formation of a ternary complex between FldIH₂–PFL-AE–AdoMet, with subsequent quaternary-complex formation between PFL-AE_red–AdoMet–pyruvate-bound PFL. Alternatively, all three proteins and the respective cosubstrates might form a single activation complex.

Radical Reduction by AdhE.

The enzyme demonstrated to catalyze radical "quenching" and conversion of PFL^• back to the nonradical, oxygen-stable form is AdhE (102, 104). This is a trifunctional protein requiring NAD⁺, CoA, and Fe²⁺ as cofactors. NAD⁺ is reduced to NADH during the deactivation reaction, and CoA also has a catalytic role, although it appears unchanged at the end of the reaction (113). In analogy to the radical chemistry involving thioester transfer in the PFL reaction mechanism, Knappe and Wagner (113) have suggested that NAD⁺ and AdhE generate a transient thiyl radical of CoA, which quenches the Cys419 thiyl radical by forming a disulfide intermediate. Many questions remain open regarding this unusual reaction, in particular, where the electron required for the reduction of the radical originates in vivo. It is likely that the buffer is the source of reductant in the in vitro reaction.

Pyruvate and NADH are strong inhibitors of radical reduction of active PFL. However, a redox poise of ≥100 mV, which equates to an NADH/NAD⁺ ratio of 0.2 to 0.4 (2), is already sufficient to induce PFL deactivation (113).

The AdhE protein is present in anaerobic cells as an oligomeric protein comprising 20 to 60 protomers and is assembled into helical rods 120 nm in length (103). It will be intriguing to determine how this large protein complex interacts with PFL^• to gain access to the radical.

Physiological Control of the Posttranslational Cycle: an NAD+-NADH Checkpoint?

Pyruvate is one of the central intermediates in cellular metabolism. Controlling its metabolic fate determines biosynthetic routes and energy metabolism and is linked to redox-responsive transcriptional regulatory networks (see below). Pyruvate metabolism is inextricably linked to the NADH/NAD⁺ ratio, and consequently to cellular redox poise.

The PFL interconversion cycle is poised to maintain the radical only when the dioxygen concentration is below a certain threshold. The activation reaction could be controlled by the available concentration of FldIH₂, which is determined by the levels of NADPH and pyruvate (Fig. 2). Radical reduction is controlled by the cellular pyruvate and NADH concentrations. If the NADH concentration is high, AdhE is active as an acetyl-CoA reductase, regenerating NAD⁺. As the NAD⁺ levels increase, it switches to a PFL^• reductase. The NADH/NAD⁺ ratio varies over an order of magnitude between aerobic growth and fermentative growth (2). The PFL-activating and -deactivating enzymes provide a monitor of this ratio and of the intracellular pyruvate concentration.

Control of the PFL Interconversion Cycle: Transcriptional Regulation.

PFL is already present at significant levels in aerobic cells (0.2% of total cellular protein), but exclusively in the deactivated form. This allows immediate activation of the enzyme by constitutively synthesized PFL-AE when the cells become microaerobic or anaerobic and consequently enables a smooth transition between pyruvate cleavage mechanisms. At low oxygen concentrations, the expression of the pfl gene is induced 12- to 15-fold, and this transcriptional regulation is mediated in concert by the redox-responsive transcription factors ArcA and FNR (51, 52, 98, 99, 100, 189, 192, 193, 195, 196, 197). The pfl gene is cotranscribed with the focA gene, which encodes a formate transport protein (215) (see Chapter Anaerobic Formate and Hydrogen Metabolism). Transcription of the operon is controlled by multiple coordinately regulated promoters (Fig. 6), which are controlled by a complex upstream regulatory region. Three promoters are located upstream of the focA gene, and there are five transcripts whose 5' ends map within the focA gene or in the focA-pfl intergenic region (Fig. 6). This transcriptional complexity is unusual for a prokaryote but has the consequence that FocA and PFL syntheses are tightly coordinated, yet it still allows the adjustment of differential cellular protein concentrations.

Fig. 6Factors affecting anaerobic regulation of the focApfl operon. The genes encoding the formate transport channel ( focA), PFL( pfl), and PFL-AE ( act) are represented by colored cylinders. The bent arrows indicate the 5' ends of transcripts, while the straight arrows indicate positive (+) or negative (−) effects on transcription of the genes by the respective transcription factors or effectors.

The regulatory region of the operon also binds integration host factor, which has an architectural role in promoting protein-protein interaction in transcriptional activation (204), and the NarL and/or NarP transcription factor, which is the regulatory component of the dual nitrate-nitrite-sensing NarLX and NarPQ two-component systems (Fig. 6) (98). Transcriptional control of operon expression is therefore intimately linked with the metabolic and redox status of the cell.

Immediately downstream of the focApfl operon is the act gene, encoding PFL-AE (184). Expression of act is constitutive and independent of focApfl expression (Fig. 6) (188). There is clearly selective pressure to keep the genes encoding PFL and its activating enzyme linked so that during lateral gene transfer both functions are acquired (239).

Expression of the adhE gene encoding the deactivating enzyme is also induced by anaerobiosis, but a mechanism different from that described for pfl is involved (see below).

Other Glycyl Radical Enzymes

Apart from PFL, the E. coli genome encodes five other confirmed or putative glycyl radical enzymes. Three of these apparently have an associated specific activating enzyme, which is encoded by an adjacent gene, while the other two are activated by PFL-AE.

The class III anaerobic RNR is encoded by the nrdDG genes and is widespread among facultative and strict anaerobes (63, 134, 213, 214). The enzyme is an α ₂ β ₂ with the α-subunit (encoded by nrdD) representing the actual catalytically active glycyl radical enzyme and the β-subunit (encoded by nrdG) representing the activating enzyme orthologue.

The E. coli genome encodes two putative PFL-like proteins, called PFLD (P32674) and PFLF (P75793), which have only a single cysteinyl residue in their active sites and therefore presumably catalyze reactions that are not related to ketoacid cleavage. Putative activating enzymes are encoded by genes that are adjacent to the genes encoding the respective cognate glycyl radical enzyme (190, 198) (Fig. 5). The physiological functions of these PFL-like proteins remain to be elucidated; however, they show some similarity to the glycyl radical enzyme benzylsuccinate synthase, which catalyzes the irreversible synthesis of benzylsuccinate from toluene and fumarate. The enzyme catalyzes the first step in the anaerobic degradation pathway of toluene (129).

TdcE.

The tdcE gene is included within a heptacistronic operon that codes for components of an anaerobically induced pathway, allowing E. coli to generate ATP from the degradation of L-threonine or L-serine (79). TdcE is highly similar at the primary structural level to PFL, and when overproduced, it can substitute for PFL in the anaerobic E. coli cell (194). The product of L-threonine dehydration is 2-ketobutyrate, which TdcE cleaves effectively to formate and propionyl-CoA. Interestingly, the structural analysis of PFL revealed that the methyl group of pyruvate can form van der Waal's interactions with Ala272, and the corresponding Ala residue in TdcE is replaced by Gly. A further difference at the active site is that in TdcE Tyr323 is replaced by a Phe residue. These changes allow accommodation of the bulkier 2-ketobutyrate in the active site (9).

Unlike expression of the focApfl operon, the tdc operon, although anaerobically induced, is repressed by glucose via the cAMP receptor protein (245). Anaerobic activation of tdc operon expression is complex and requires the divergently transcribed tdcR gene product and the product of the first gene, TdcA (191). TdcA is a LysR-like transcriptional regulator, and although the mechanism of anaerobic regulation is not fully understood, it is thought that TdcA might interact with a metabolite that accumulates during fermentative growth (191).

YfiD.

YfiD (P33633) belongs to a family of proteins found in many facultative anaerobes that also have a PFL. Homologues are also found in several bacteriophages, for example, phage T4. YfiD has a mass of 14.3 kDa, and its N-terminal 67 amino acids have no similarity to those of any known protein, while the C-terminal 60 amino acids are very similar to the C-terminal 60 amino acids of PFL (113). Remarkably, YfiD can restore activity to PFL molecules that have been inactivated through exposure to dioxygen (233). Therefore, the protein is thought to represent a glycyl radical module that can associate with glycyl radical enzymes damaged by oxidative stress. Indeed, association with YfiD also reconstituted wild-type activity in a truncated PFL derivative devoid of the C-terminal 26 amino acids (including Gly734 [233]). It has also been shown that association of YfiD with PFL stabilizes the glycyl radical in YfiD and significantly reduces the K_m for PFL-AE, which introduces the radical (233, 246).

Expression of the yfiD gene is induced by acid stress and is optimal in the microaerobic range, which correlates well with recent findings for pfl expression (15, 70, 141). A yfiD knockout mutant has a severely altered spectrum of fermentation products, with increased formate levels relative to those of acetate and ethanol, suggesting reduced FHL activity (246). Furthermore, pyruvate, D-lactate, and succinate are the main excreted products, suggesting an inhibition of pyruvate cleavage. It has been argued, therefore, that the function of YfiD is to repair dioxygen-damaged glycyl radical enzymes to allow maintenance of metabolism until de novo synthesis can provide new enzyme (113). This hypothesis is particularly pertinent in the case of PFL, which produces formate, serving as the reductant for ribonucleotide reduction by activation of the other glycyl radical enzyme, class III RNR. It is perhaps notable that strict anaerobes lack YfiD-like proteins.

The proposed role of YfiD as a glycyl radical module raises several important questions. Can YfiD "rescue" other oxygenolytically damaged glycyl radical enzymes, and if so, how is specificity conferred? How does YfiD recognize that an enzyme has become damaged? Perhaps most significant of all, how does YfiD interact with PFL to abstract the hydrogen atom from Cys419? In this regard, it is perhaps significant that in all members of the family examined so far, the glycyl residue forming the radical is located in the C-terminal 20 to 30 amino acids of the protein (113).

ACETYL-CoA METABOLISM

PTA and ACK

During fermentation, acetyl-CoA may be converted to either ethanol or acetate. In practice, a mixture is made, and the ratio depends on the redox level of the carbon source (see above). The first step in acetate formation is catalyzed by PTA (or acetyl-CoA–orthophosphate acetyl transferase), which converts acetyl-CoA plus inorganic phosphate to acetyl phosphate and CoA. The second step is catalyzed by ACK (or ATP-acetate phosphotransferase), which converts acetyl phosphate plus ADP to acetate plus ATP (Fig. 1). This reaction sequence therefore generates an extra ATP in addition to those derived from glycolysis. Even when growing aerobically, E. coli often excretes substantial amounts of acetate. This occurs when the carbon flux exceeds the capacity of the central metabolic pathways, in particular, the tricarboxylic acid (TCA) cycle (118). It is especially noticeable for cells growing in rich medium with plenty of glucose or other sugars. This aerobic acetate excretion also relies on the PTA/ACK pathway.

Uptake of Acetate.

Both the PTA and ACK reactions are freely reversible. Indeed, the PTA and ACK reaction sequence can be used by E. coli for aerobic growth with acetate as a carbon source. Aerobic uptake of acetate may also be mediated by the enzyme acetyl-CoA synthetase (ACS), which converts acetate plus ATP to acetyl-CoA plus AMP and pyrophosphate and proceeds via an enzyme-bound acetyl adenylate (118). ACS is not expressed anaerobically, and its regulation and metabolic role are discussed elsewhere. The possession of two alternative pathways means that pta/ack mutants of E. coli and Salmonella still grow aerobically on acetate. However, certain other enteric bacteria, such as Enterobacter aerogenes, apparently lack ACS and rely solely on the PTA/ACK pathway for aerobic growth on acetate (23, 24).

The mechanism for the uptake and/or excretion of acetate is poorly understood. Originally, two separate transport systems were proposed, based on kinetic evidence (234). More recently, the yjcG gene, which is cotranscribed with acs, has been shown to encode an acetate uptake carrier and has been renamed actP (66). The authors also propose the existence of a second transport system for excretion and uptake of acetate.

Biochemistry of PTA and ACK.

ACK (EC 2.7.2.1) is a protein of 400 amino acids with a molecular mass of 43,297 Da (147) and probably functions as a homodimer. The enzyme is cold labile but may be reactivated by MgATP plus acetate. This behavior resembles that of enzyme I of the phosphotransferase system (116). The K_m value of ACK for acetate is high (7 mM), whereas the K_m value of the enzyme for acetyl phosphate is in a physiologically reasonable range at 0.16 mM (61). This implies that the physiological role of the enzyme is for ATP generation rather than for acetate uptake. The V _max values of the enzyme are similar in both directions.

Purified ACK was shown to require Mg²⁺, although Mn²⁺ can substitute. Na⁺ and Li⁺ are inhibitory (61). However, a recent systematic search for E. coli proteins with zinc-binding activity has revealed that ACK binds Zn²⁺ (101). This is in accord with earlier enzymological work indicating that a variety of divalent cations (Mg²⁺, Mn²⁺, Co²⁺, Zn²⁺, and Cd²⁺) can replace each other in the ACK reaction (185), so perhaps Zn²⁺ is the actual cation used by the enzyme in vivo.

The ACK reaction appears to involve a phosphoenzyme intermediate (4, 60, 91). However, both substrates must be added before either product is released, suggesting a random sequential mechanism of the enzyme (91). The phosphoryl group is probably carried by an aspartyl or a glutamyl residue, as deduced from the response to varying the pH (60). An essential arginine is probably also involved in binding the negatively charged phosphate group (244). Stereochemical analysis has shown that the configuration at the phosphoryl group is inverted during phosphotransfer by ACK (16). The phosphoryl group may also be transferred from ACK to enzyme I of the phosphotransferase system (60). This might represent a mechanism for coupling regulation of sugar transport to the energy charge (the ratio of ATP to ADP) and/or the level of acetyl phosphate.

A second enzyme with ACK activity is known in E. coli. This is the TdcD protein, encoded by the tdc operon for anaerobic threonine degradation. TdcD presumably functions as a propionate kinase in vivo, although it will also use acetate (79).

PTA (EC 2.3.1.8) is a protein of 713 amino acids (146). The subunit structure is uncertain, although genetic studies indicate that only a single type of polypeptide is involved (23, 130). Experimental observation gives a mass between 160,000 and 250,000 Da, implying a dimer or trimer (200). The enzyme is rather unstable but has been purified after stabilization with ammonium sulfate. The K_m value for the enzyme for acetyl phosphate is 3 mM, and the K_m value for CoA is 0.3 mM (200). The K_m value for acetyl phosphate varies with the pH and drops 10-fold between pH 7.8 and 5.8 (216). The enzyme is positively allosterically regulated by pyruvate and negatively regulated by NADH, ATP, and ADP. Pyruvate activation results in a lowering of the K_m value for acetyl phosphate (216), but the V _max remains unchanged.

The 350-amino-acid N-terminal region of the PTA protein of E. coli and several other bacteria is not required for PTA activity. Sequence analysis indicates a relationship of this domain to a family of ATP-dependent amido-ligases that includes dethiobiotin synthetase and the domains of two amidotransferases involved in cobalamin biosynthesis (64).

Metabolic Effects of Mutations Affecting PTA and ACK.

Under anaerobic conditions, pta mutants grow poorly or not at all on fermentable sugars. However, double mutants defective in both pta and adhE regain the ability to grow and produce lactic acid as the major fermentation product (73). This implies that the original growth defect of the pta mutants was due to disturbance of the redox balance. Further introduction of a ppc (PEP carboxylase) mutation eliminates the small amount of succinate produced and allows production of pure D-lactate by fermenting E. coli (29).

Mutants defective in the ack or pta gene grow poorly on acetate aerobically, despite the existence of the alternative ACS route for acetate uptake. This is apparently the result of defects in ack or pta preventing induction of the acs gene (118). Since ack and pta mutants behave the same in this respect, the induction of acs cannot be dependent on acetyl phosphate. Mutants defective in the pta gene grow more slowly than wild-type strains aerobically on sugars, but little or no growth impairment is seen with gluconeogenic carbon sources, such as succinate (75). When grown on glucose, pta mutants excrete alternative by-products, such as pyruvate, D-lactate, and L-glutamate, instead of acetate (30, 249).

Engineered strains of E. coli are now being used to produce a variety of products, including recombinant human proteins for pharmaceutical purposes. One of the problems encountered is that high levels of acetate and/or low pH may inhibit expression of recombinant proteins and may damage the protein structure. Consequently, several groups have investigated the potential use of mutants defective in pta and/or ack to reduce acid production during the growth of E. coli strains harboring genes for recombinant proteins. Despite alternative by-products being produced, the overall medium acidity is significantly decreased, and improved yields of recombinant protein are generally obtained in pta mutants (255). Such mutants may be easily obtained, as shown long ago by Guest (71), by selection for fluoroacetate resistance.

Regulation of PTA and ACK.

Synthesis of both PTA and ACK enzymes is largely constitutive, and results of mutant studies suggest that the ackA and pta genes encoding the two enzymes form an operon that is located at 52.0 min on the chromosomes of both E. coli and Salmonella (23, 71, 119, 130, 146, 147, 171). Earlier reports indicated that expression of the ackA-pta operon is unaffected by acetate, formate, and pyruvate and is only slightly increased by anaerobic conditions (119).

More recent data suggest that the levels of PTA and ACK do change under certain conditions; however, the changes are relatively small. When cells are grown on acetate, PTA and ACK levels decrease ~2-fold (166, 167). Acs levels increase nearly 10-fold under these conditions, implying that this is the major route for acetate uptake (167). Addition of acetate to cells growing in rich broth also depresses PTA levels (106). When E. coli is grown on glycerol, ackA is significantly down-regulated (172). On such a carbon source, growth is slower than on glucose, and there would be little or no surplus acetyl-CoA to be excreted as acetate instead of being metabolized by the TCA cycle. Indeed, TCA cycle enzymes are up-regulated in cells grown on glycerol (172).

A known system that regulates the ack-pta operon is the CreBC two-component system, which was discovered because the sensor compound CreC (originally called PhoM) was able to phosphorylate PhoB in strains lacking PhoR, conferring some residual phosphate regulation on these strains (238). Although the physiological signal that activates CreBC remains elusive, it has been shown that the ackA-pta operon is regulated by CreBC (7). Expression of the ackA-pta operon and several other cre regulon genes is induced via CreBC during growth in glucose minimal medium. Insertional inactivation of creC abolishes induction, and conversely, constitutive creC mutants cause higher expression under noninducing conditions (7).

Some time ago, Pascal et al. (171) described a second gene, ackB, affecting ACK activity. Although no further reports have been published, the original ackB mutant does indeed lack ACK, and the defect is not cotransducible with the ackA-pta operon (D. P. Clark, unpublished data).

Regulatory Effects of Acetyl Phosphate.

Although the ACK/PTA reaction sequence responds only slightly to regulation, several regulatory roles have been ascribed to the metabolic intermediate acetyl phosphate. Acetyl phosphate has been shown to phosphorylate the regulator proteins of several two-component regulatory systems. Usually this occurs only in the absence of the cognate sensor protein, suggesting that it may be of little in vivo significance. For example, the regulator of the phosphate regulon in E. coli, PhoB, is phosphorylated by acetyl phosphate in the absence of the cognate or alternative sensory component, PhoR or CreC (105, 238). The osmoregulator OmpR is also phosphorylated and modulated in its activity by low-molecular-weight phosphate donors, such as acetyl phosphate (78) (see below). Additionally, acetyl-phosphate also seems to play a role in protecting cells against carbon starvation by an as yet unknown mechanism (163).

Although transcription of the ompF porin gene is dependent upon acetyl phosphate only when EnvZ is absent (88, 125), the OmpC level has been shown to increase as acetyl phosphate accumulates, even in the presence of EnvZ (151, 201). Certain variants of the OmpR protein with single amino acid substitutions can no longer be phosphorylated by acetyl phosphate but retain the ability to be phosphorylated by EnvZ (148, 225). For example, the V203M variant of OmpR constitutively activates ompF but fails to express ompC (225), whereas the T83I variant fails to activate either ompF or ompC (148).

Variants of the converse type to those described above have been found for the PhoP response regulator that controls virulence gene expression in S. enterica (28). A PhoP species with the single amino acid substitution S93N was able to function in the absence of its cognate sensor (PhoQ). The mutant PhoP was more efficiently phosphorylated by acetyl phosphate.

Despite these sophisticated analyses, it is still unclear in many cases whether the effects of acetyl phosphate are physiologically relevant or, in some cases, are even due to acetyl phosphate. However, Heyde et al. (80) found that the external pH affected the acetyl phosphate level, which in turn modulated OmpR phosphorylation and led to pH-dependent changes in porin expression.

Acetyl phosphate has also been implicated in the control of flagellar rotation by exogenous acetate (44). However, later work implicated both ACK and ACS as mediators of the response to acetate (8, 177). Here, acetylation of the response regulator CheY (rather than phosphorylation) is involved. To add to the complexity, acetyl phosphate has been shown to affect synthesis of the flagellar apparatus (176). As for the effects on regulation of the porins, this proves to involve phosphorylation of OmpR by acetyl phosphate. Phosphorylated OmpR, generated either by EnvZ in response to a change in osmolarity or by acetyl phosphate, negatively regulates the expression of flagella via the flagellar master operon regulator FlhDC (201).

The accumulation of acetyl phosphate is diagnostic of the incoming carbon flux being too great to be accommodated by the TCA cycle. In other words, the cells are well fed and do not need to forage for more nutrients. Thus, from a physiological viewpoint, acetyl phosphate is a metabolite well suited to down-regulate flagellar motion and to modulate porin expression, shifting synthesis of the porin isoforms to those with smaller porin size.

The scenario of acetyl phosphate working via two-component regulatory systems has been complicated by other observations using pta and ackA mutants. Effects on ompC and ompF expression that were independent of OmpR have been observed. It was suggested that the stress-induced sigma factor (RpoS) system might be involved (132). The RpoS regulon is induced by added exogenous acetate and is also expressed at higher levels in an ackA-pta deletion mutant (106). A possible mechanism is that an excess of acetyl-CoA is responsible for regulation in these cases. Indeed, blocking the PTA/ACK pathway does result in accumulation of acetyl-CoA (227). Furthermore, it has been shown that exogenous acetate represses PhoPQ signaling via internal build up of acetyl-CoA, which then inhibits the autophosphorylation of PhoQ (127).

Therefore, when assessing the effects of ackA-pta mutations on gene expression, the following factors should be taken into account:

Blocking the PTA/ACK pathway decreases acetate efflux into the medium. Consequently, less acidification of the medium occurs.
Blocking the PTA/ACK pathway reduces acetyl-CoA metabolism, and the level of internal acetyl-CoA therefore increases.
Pyruvate accumulation may occur and result in the formation of other fermentation products, such as D-lactate. This in turn may trigger the ArcAB system (65).
Mutants lacking both ACK and PTA will be devoid of acetyl phosphate under all conditions. Those lacking one or the other enzyme will show a lack or a surplus of acetyl phosphate, depending on the growth conditions (i.e., in which direction the PTA/ACK pathway is running under the prevailing physiological conditions).
Finally, it is possible that some of the observed regulatory effects are due to the N-terminal amidoligase-related domain of the PTA protein, whose role is presently unknown (64).

AdhE and the Formation of Ethanol.

In E. coli, the reduction of acetyl-CoA via acetaldehyde to ethanol is carried out in two steps by a single protein, known as AdhE (43, 67, 187). Each step reoxidizes an NADH molecule (Fig. 1). Both reactions are reversible, but in wild-type E. coli, the AdhE protein is expressed only under anaerobic conditions, and its physiological role is the conversion of acetyl-CoA to ethanol (36, 37, 48, 187). In addition, the enzyme specifically quenches the free radical of PFL and functions as a PFL deactivase (see above) (102, 104).

Since the PTA/ACK pathway is largely constitutive, the regulation of carbon flow is mainly due to alterations in the level of AdhE. In particular, expression of the adhE gene responds to the NADH levels inside the cell (see below).

The Multifunctional AdhE Protein.

Initially, it was believed that the AdhE and acetaldehyde dehydrogenase (ACDH) reactions were catalyzed by distinct enzymes in E. coli, as they indeed are in many organisms. However, both genetic and biochemical analyses have clearly demonstrated that the AdhE protein catalyzes both reactions (67, 102). Sequence homology indicates that the AdhE protein consists of an N-terminal (CoA-acylating) ACDH domain and a C-terminal alcohol dehydrogenase (ADH) domain (Fig. 7). In addition, AdhE also shows PFL deactivase activity, probably located in the N-terminal domain (102, 104). All three functions of the enzyme involve redox processes and are highly dependent on ferrous iron and NAD(H) (102, 104).

Fig. 7Proposed domain organization of AdhE of E.coli. The proteolytic cleavage site described in reference 102 is located at amino acid position 762. The resulting 86-kDa truncated product retains AHD activity and, based on amino acid sequence analysis, is localized to amino acids 450 to 772.

In its PFL-deactivase role, AdhE catalyzes the removal of the glycyl radical from PFL, and possibly also the TdcE enzyme (79), but it does not affect the alternative acid-inducible YfiD protein (246). The adhE gene from Lactococcus lactis was cloned by functional complementation using an E. coli adhE mutant. L. lactis also possesses PFL, but this is not subject to deactivation by its AdhE protein (152). Several other industrially important lactic acid bacteria were also shown to possess adhE orthologues. Although they are typically regarded as homolactate fermenters, these organisms sometimes show mixed acid fermentation on sugars other than glucose (152).

The AdhE protein consists of 891 amino acid residues and is one of the largest polypeptides found in E. coli. AdhE forms a homopolymer comprising between 20 and 60 identical subunits. As each subunit has a molecular mass of 96,000 Da, this yields a mass of several million Da for the native protein (102, 104). Electron microscopy identified the ADH protein as a left-handed helical rod ranging from 45 to 120 nm in length, most frequently with a length of 80 nm. This finding immediately suggested identity between AdhE and the enigmatic helical structures ("spirosomes") previously identified by electron microscopy in many anaerobic and facultative bacteria, including E. coli (144). Moreover, the findings are in accord with the observation that ACDH activity of E. coli is associated with a large soluble protein particle (199). Incubation of purified AdhE with NAD⁺ and Fe²⁺ ions causes the protein structure to expand along its helical axis from 80 nm to a length of 130 nm (102). This implies that a major conformational change occurs within the polymeric enzyme that is probably associated with a shift of its catalytic functions.

As already mentioned, all three activities of AdhE require Fe²⁺ ions, but the protein must also be able to interact with several substrates. The substrates of the PFL deactivase reaction are radical-bearing PFL and NAD⁺, plus stoichiometric amounts of CoA. The ADH activity uses acetaldehyde plus NADH or ethanol plus NAD⁺ as substrates; ACDH uses acetyl-CoA and NADH or acetaldehyde, coenzyme A,and NAD⁺ as substrates (102). Where these different activities are accommodated on this remarkable protein remains to be fully elucidated experimentally. Sequence analysis shows only a single recognizable NAD⁺-binding site, located between the ACDH and ADH domains of AdhE, implying that the site may be shared by the two enzyme activities. Support for this comes from kinetic measurements showing that the K_m for NAD⁺ (32 μM) is identical for the ACDH and ADH reactions (J. T. Koepke and D. P. Clark, unpublished data). Furthermore, the two reactions showed identical K_i values for several different triazine dyes that are potent competitive inhibitors, acting as NAD⁺ analogues.

Nonetheless, Kessler and colleagues have been able to separate ADH activity physically from the other two enzyme activities by cleaving the enzyme into an 86- and a 14-kDa polypeptide by controlled treatment with chymotrypsin (102). The larger peptide retained ADH activity but lost ACDH and PFL deactivase activities. Taking into account sequence similarities with ADHs from other sources, this localizes the ADH activity to a region spanning amino acids 450 to 772, still present in the large fragment (102). ACDH and PFL deactivase activities of AdhE have in common their requirement for CoA and loss of activity after chymotrypsin treatment. The latter observation also suggests that both the N-terminal amino acids 1 to 449 and the C-terminal region (amino acids 763 to 891) of AdhE are necessary for these activities (Fig. 7). It is probable that either one or both of these regions also will harbor the polymerization domain of the enzyme (102).

The N-terminal ACDH domain of AdhE is homologous to a large family of aldehyde dehydrogenases that are widespread in organisms. Most of these convert acetaldehyde to acetate. However, a small subgroup (the CoA-acylating ACDHs [EC 1.2.1.10]) converts acetaldehyde to acetyl-CoA and aldehydes to the corresponding acyl-CoA thioesters (140, 169, 202, 205). The location of the CoA-binding site in these enzymes is uncertain. One possibility is that it is shared with the NAD⁺-binding site, as suggested by the fact that NADH dissociates before CoA binds (140, 202).

Isoenzymes of AdhE (alcohol NAD⁺ oxidoreductase [EC 1.1.1.1]) are widespread in nature and play key roles in both fermentative ethanol formation and the consumption of alcohol as a carbon source under aerobic conditions. Several ADH families that use NAD⁺ as a coenzyme are known. The most prevalent comprise a superfamily of zinc metalloenzymes ("long-chain ADHs"), often of broad specificity and capable of oxidizing a wide range of aliphatic and aromatic alcohols to the corresponding aldehydes and ketones, and a second superfamily of metal-dependent "short-chain ADHs," which are often involved in β-oxidation pathways (180). However, the ADH domain of the AdhE protein of E. coli belongs to a smaller family of ADHs that have Fe²⁺ as their active-site metal and, apart from possession of an NAD⁺-binding motif, are unrelated in sequence to the other ADH superfamilies (67). Fe-ADHs belonging to this group may occur as monofunctional proteins. For example, ADH III of Zymomonas mobilis (41) and ADH IV of Saccharomyces cerevisiae (241) exhibit strong similarity to just the portion of the E. coli AdhE polypeptide (amino acids 450 to 850 [Fig. 7]) that has been correlated with its ADH activity. Alternatively, some Fe-ADHs are similar to the C-terminal domains of AdhE-like proteins. The members of the Fe-ADH group are typically sensitive to oxygen inactivation and are mostly found in obligate or facultative anaerobes.

Taxonomic Distribution of AdhE-Like Proteins.

Several multifunctional two-domain AdhE-like proteins are known in addition to that of E. coli. These dual-domain AdhE proteins may be the result of the evolutionary fusion of separate genes encoding a CoA-linked ACDH and an Fe-activated AdhE. They are found in a variety of bacteria, including facultative anaerobes, obligate anaerobes, and cyanobacteria, and also in some primitive protozoa that live by fermentation.

The AdhE protein of Clostridium acetobutylicum is part of an operon for butanol production and displays butanol- and CoA-linked butyraldehyde dehydrogenase activities (58, 159). This is interesting with regard to the finding that the AdhE of serovar Typhimurium works well with butanol and longer alcohols up to about C₈, whereas the E. coli enzyme is poorly active beyond C₄ (46). The Salmonella enzyme also has a lower K_m value for the corresponding alcohol substrates than AdhE from E. coli, although the K_m value for NAD⁺ is similar. The substrate chain length specificities of other putative multifunctional AdhE proteins are as yet unknown.

Although orthologues of AdhE are widespread, use of a multifunctional protein is not the rule for all anaerobic bacteria. Clostridium kluyveri, for example, possesses two distinct proteins: an AdhE of the Fe-Adh class and a CoA-linked ACDH. However, inside the cell they form a complex that operates like the single AdhE protein found in E. coli (205).

The protozoans Entamoeba histolytica (25, 248) and Giardia lamblia (186) also possess adhE genes with products having ~50 to 60% amino acid identity to that of E. coli. This has suggested that lateral transfer of fermentation genes from prokaryotes to intestinal protozoa has occurred (186). Indeed, the AdhE of Giardia is closer to that of E. coli (49% identity) than it is to that from Entamoeba (43% identity). Moreover, the Entamoeba AdhE can functionally replace the E. coli enzyme and restore anaerobic growth to an E. coli adhE deletion mutant (252), and it even forms helical rods resembling those of E. coli (6, 25).

Genetics of the AdhE Protein.

Several screening and selection procedures have been used to isolate mutants with altered AdhE levels. These approaches included screening for mutants capable of aerobic growth on ethanol as a sole carbon source, yielding AdhE-overproducing strains (36, 37); rescreening for loss of growth on ethanol of such AdhE-overproducing mutants (36, 243); and selecting mutants resistant to suicide substrates, such as chloroethanol, chloroacetaldehyde (43), or allyl alcohol (136). Even screening for loss of the ability to reduce the redox dye benzyl viologen, which is normally used to isolate hydrogenase-defective or FHL-defective mutants, resulted in a few AdhE-negative mutants (27). The genetic link between ADH and ACDH activities (36) was already evident in early experiments, since all mutants lacking ADH enzyme activity were also defective in ACDH activity. Moreover, the mutants were unable to ferment any hexoses or sugar alcohols and grew anaerobically either by fermenting the more oxidized glucuronate or by anaerobic respiration with fumarate or nitrate as an electron acceptor (27, 36, 37, 73, 104).

Several temperature-sensitive mutants in which both ADH and ACDH activities were thermolabile were isolated, and mutants with increased ADH activity also had concomitantly increased ACDH activity, again implying that both are activities of a single protein (36, 136).

Defects in the acd gene (which is probably allelic to ubiH, coding for a 2-octaprenyl-6-methoxyphenol 4-monoxygenase involved in ubiquinone biosynthesis) result in the inactivation of the ACDH activity of the AdhE protein, although the ADH activity is unaffected (37). Activity is restored by adding ubiquinone-0 to enzyme preparations. Abolition of ubiquinone synthesis by defects in other ubi genes had no effect, suggesting that the ACDH activity is specifically inhibited by phenolic intermediates of ubiquinone synthesis that accumulate in acd/ubiH mutants (74).

Transcriptional Regulation of adhE Expression.

Expression of the adhE gene is induced by anaerobiosis (11, 123). However, this induction is independent of the Fnr and ArcA transcription factors (33, 123). Anaerobic induction appears to be largely due to the accumulation of reduced NADH rather than an effect of oxygen per se. Nitrate represses adhE expression largely via the NarXL two-component regulatory system, with NarL acting as a repressor of adhE transcription (33, 123). In addition, nitrate respiration alters the cellular redox potential, exerting an indirect effect on adhE (11, 33).

There appears to be a direct correlation between the NADH/NAD⁺ ratio and enzyme synthesis; the higher the ratio, the more AdhE is synthesized (123, 124). Consequently, AdhE levels are higher under anaerobic conditions. Indeed, the NADH/NAD⁺ ratio increases as the oxygen concentration in the medium decreases, and this correlates well with the amount of ethanol generated by fermentation (3). In addition, AdhE levels are higher in cells growing anaerobically on sugar alcohols than in those growing on hexoses (123). This is also true of AdhE levels in Salmonella spp. (46). The AdhE protein itself apparently affects the regulation of its own gene, since gene expression is dramatically enhanced in an adhE mutant (46). Introduction of mutant adhE alleles producing high levels of inactive AdhE protein gave results equivalent to those seen in the absence of the AdhE protein (124). This indicates that the effect of AdhE is indirect and is probably due to the accumulation of high NADH levels in cells lacking functional AdhE, which are consequently unable to dispose of NADH by making ethanol.

The adhR (yhdY) gene, located at 3,419 kb, is known to regulate adhE (located at 1,297.3 kb) expression (39, 124). However, interpretation of the role of adhR was originally complicated by the presence of additional mutations, some of which were in the nearby rng gene (see below). Defects in adhR (in the absence of other mutations) greatly decreased expression of adhE, implying that AdhR acts as an activator for adhE transcription (45). Furthermore, the AdhR protein binds to an adhE promoter fragment in an NAD(H)-dependent manner. Thus, the adhR gene product appears to mediate the previously described NADH-dependent induction of the adhE gene (123, 124).

Expression of adhE is also repressed by the Cra (formerly called FruR) regulator protein, which is involved in the cAMP-independent modulation of a number of operons involved in carbon catabolism (157). Cra was shown to be displaced from the adhE DNA by its effector-coregulator, fructose-1-phosphate (157). Some mutations causing aerobic overproduction of AdhE (previously called adhC ) were shown to carry a point mutation in the Cra-binding site of the adhE promoter (95). Since Cra responds to metabolite levels rather than to aeration or redox conditions, it seems likely that the physiological role of Cra is to tighten the control of adhE transcription under aerobic conditions.

Recent use of microarray screening techniques has confirmed that adhE is expressed under microaerobic or anaerobic conditions (172). In addition, adhE was down-regulated during aerobic growth on acetate (relative to control cultures on other carbon sources) (166). This could be due to lower NADH levels in cells growing on acetate and/or to repression by Cra in response to altered levels of fructose phosphate.

Transcription of adhE requires the Fis protein as an accessory factor (154). The adhE upstream region appears to contain three transcriptional start sites (45, 155). The first promoter, starting at −188 bp relative to the start of the adhE coding sequence, appears to function only in experimental constructs lacking upstream sequences and not under physiological conditions. The promoter is activated by Fnr and RpoS, and the transcript formed does not require RNase III processing (155). The second promoter, at −292 bp relative to the translation initiation codon, appears to be the major physiological promoter (45). However, there are conflicting claims that this promoter is either activated by RpoS (155) or down-regulated in the presence of acetate, which induces the RpoS regulon (166). The third, and most distal, promoter is poorly characterized but appears to function when glucose is the carbon source (45, 122).

Regulation of adhE Expression by RNA Processing.

The adhE gene is regulated not only at the level of transcription but also at the posttranscriptional level. Both translation and degradation of the adhE mRNA are regulated. When first transcribed, adhE pre-mRNA has a long 5' leader sequence that apparently folds into a secondary structure, preventing access to the ribosome-binding site (5). Translation of the adhE transcript is initiated only after this leader sequence has been cleaved off by RNase III (formerly RNase C), the product of the rnc gene. Consequently, defects in the rnc gene prevent expression of adhE at the level of translation, and rnc mutants do not grow anaerobically by glucose fermentation, although they can still grow on more oxidized substrates, such as glucuronate (5).

Degradation of matured adhE mRNA involves RNase G, encoded by the rng gene (previously known as cafA). In rng knockout mutants, the level of AdhE protein is much higher and the half-life of adhE mRNA is 2.5 times longer than in wild-type cells (226). RNase G is involved in rRNA processing and also in the turnover of the mRNAs of a very small number of metabolic genes, in particular adhE and eno (encoding enolase) (96). A mutant isolated long ago (39) for increased expression of adhE was recently shown to be mutated in rng. The rng430 mutation has a single amino acid alteration, Gly341Ser, which results in an altered RNase G that retains the ability to process 16S rRNA but no longer degrades adhE mRNA (230). Since the effects of RNase G on adhE are posttranscriptional, they depend on the availability of adhE mRNA and are thus most noticeable under conditions where the adhE gene is transcriptionally derepressed (38).

Oxygen Sensitivity of the AdhE Protein.

Enzymes of the Fe-ADH family are highly sensitive to oxygen inactivation. Thus, wild-type AdhE protein is rapidly inactivated on exposure to air by Fe-catalyzed oxidation. However, constitutive mutants ("adhC") that grow aerobically on ethanol as a carbon source are known (36). They not only exhibit alterations in the promoter sequence, allowing aerobic expression of the adhE gene, but also have alterations in the AdhE protein that make it more resistant to oxidative damage (85). This aerotolerant form of AdhE has two amino acid changes, Ala267Thr and Glu568Lys. However, the Glu568Lys substitution is sufficient to protect the enzyme against oxidation. Furthermore, replacement of Glu568 by virtually any amino acid also resulted in AdhE isoforms that were active under both aerobic and anaerobic conditions (85, 153). A trade-off of the aerotolerant AdhE was a noticeably higher thermosensitivity than the wild-type protein.

Curiously, aerobic growth on ethanol also required the presence of the chaperone DnaK (a member of the Hsp70 family), which appeared to protect the mutant AdhE (Ala267Thr; Glu568Lys) against metal-catalyzed oxidation (53). Recently, Echave et al. (54) have suggested that AdhE itself plays a role in protection against oxidative stress, despite being anaerobically induced. They claim that cells with the adhE gene deleted could not grow aerobically in minimal media. However, many strains of E. coli carrying both deletion and insertion mutations in adhE have been grown in the laboratory of one of the authors (Clark, unpublished), and all of them grew perfectly well aerobically in minimal media.

Choice between Different Fermentation Pathways.

E. coli makes two major choices between alternative fermentation schemes: lactate fermentation versus ethanol-acetate fermentation and ethanol versus succinate formation (Fig. 1). At low pH, the cells tend to generate lactate rather than a mixture of ethanol plus acetate plus formate. As already discussed, the ldhA-encoded D-LDH is induced at low pH under anaerobic conditions (Fig. 8) (145). Furthermore, it seems that lactate fermentation may be favored under low-iron conditions (see above). Fermentation product compositions have usually been compared using cultures grown in minimal medium and with glucose as a carbon source. Consequently, for many years it was not realized that in rich medium and in the absence of glucose E. coli generates large amounts of succinate rather than ethanol (31, 50). Abolition of catabolite repression by mutations in ptsG, encoding enzyme II of the glucose phosphotransferase system, or growth on non-catabolite-repressing sugars results in a switch in the major reduced fermentation product from ethanol to succinate, but only in cultures in rich medium. At present, it is unclear how this metabolic switchover is regulated (Fig. 8).

Fig. 8Summary of effects of environmental and metabolic signals on regulation of synthesis and activity of fermentative enzymes. The large pluses next to green arrows indicate positive effects of the metabolite, and the minus next to the red arrow indicates anaerobiosis. Fermentation products are shown in green.

A final observation concerns the effect of heavy water on the distribution of fermentation products (20). Increasing levels of D₂O result in a decline in the production of ethanol plus acetate, whereas lactate synthesis is relatively unaffected. Higher levels of D₂O have also been shown to increase the rate of carboxylation of phosphoenol pyruvate to oxaloacetate (83). This would be expected to divert flux to succinate production and away from ethanol synthesis. However, the regulatory mechanisms involved in this metabolic transcription are totally obscure.

Other AdhEs of Enteric Bacteria.

Some mutants of Salmonella that have been selected for aerobic growth on ethanol differ from the corresponding mutants of E. coli (36) in showing no alterations in the level or oxidation resistance of AdhE (47). Instead, these adhI mutants expressed very high levels of a novel ADH, named AdhI because of its high activity with isopropanol. Unlike AdhE, this enzyme does not appear to have an associated ACDH activity. Both the parental Salmonella strain and alcohol-utilizing adhI mutant synthesize an aldehyde dehydrogenase that presumably catalyzes the oxidation of acetaldehyde to acetate in the ethanol catabolic pathway (47). The physiological role of AdhI is unknown.

Two other adh gene loci are present in the E. coli genome. Both genes are identified solely based on sequence similarity of the derived products, and none of the putative enzymes has been characterized. Despite prior use of the term adhC to refer to constitutive mutants affecting adhE expression (36), the locus encoding a glutathione-dependent formaldehyde dehydrogenase (a class III AdhE) at 8.1 min has confusingly been named adhC. The adhP locus at 33.4 min encodes a presumptive propanol-utilizing ADH.

Formation of 2,3-Butanediol

The 2,3-Butanediol Pathway.

In 1906, Harden and Walpole (77) found that the fermentation product pattern of Aerobacter aerogenes (now E. aerogenes) differs from that of E. coli due to the presence of neutral compounds, later identified as 2,3-butanediol and acetoin (acetylmethylcarbinol). The capacity to produce 2,3-butanediol is widely distributed among bacteria. Among the enteric bacteria, Klebsiella, Enterobacter, and Serratia make significant levels of butanediol. Other examples are found in the genera Bacillus, Lactobacillus, and Aeromonas. In all cases, major products from other fermentation pathways accompany 2,3-butanediol, in addition to the small amount of acetoin made by the 2,3-butanediol pathway itself. In the enteric bacteria, these other products typically include ethanol and formate (137).

Table 3 compares the fermentation balance of Klebsiella oxytoca (previously Aerobacter indologenes) (182) with that of E. coli (14). E. coli produces no 2,3-butanediol or acetoin, whereas butanediol is one of the major products of K. oxytoca. Three enzymes are involved in the conversion of two molecules of pyruvic acid into one molecule of 2,3-butanediol (Fig. 9): α-acetolactate synthase (ALS), α-acetolactate decarboxylase (ALDC), and acetoin reductase (AR).

Fig. 9Pathway of 2,3-butanediol formation from pyuvate by Enterobacter and Klebsiella spp. AHAS, anabolic α-acetohydroxyacid synthase; 2-OA, 2-oxo-acid.

Table 3Comparison of fermentation products of E.coli and K.oxytoca

The widely used Voges-Proskauer test is used to identify bacteria based on possession of the butanediol pathway (55). This test actually detects acetoin, which is usually present in much smaller amounts than butanediol. Acetoin is converted to diacetyl (2,3-butanedione) by reaction with oxygen in the presence of α-naphthol (Fig. 9). The diacetyl then reacts with the guanidino groups of compounds such as arginine, which is present in the nutrient broth. This yields the typical red color indicating a positive result.

ALS.

ALS (EC 4.1.3.18) contains the cofactor thiamine pyrophosphate (TPP). It converts two molecules of pyruvate to α-acetolactate plus carbon dioxide. The first molecule of pyruvate forms an adduct with the TPP and is decarboxylated, giving a hydroxyethyl-TPP intermediate. The resonance-stabilized carbanion then attacks the keto group of a second molecule of pyruvate, yielding α-acetolactate (Fig. 9).

The fermentative ALS has been purified from both E. aerogenes (209) and Serratia marcescens (139). ALS is a homodimer of 58-kDa subunits (89). The fermentative or catabolic ALS must be distinguished from the biosynthetic α-acetohydroxy acid synthases involved in isoleucine and valine biosynthesis (Fig. 9). In the cases of Enterobacter and Serratia, the fermentative ALS shows a narrow pH optimum around pH 6.0 and has been described as the "pH 6.0" acetolactate-forming enzyme in contrast to the biosynthetic ALS. However, stationary-phase cells of Bacillus subtilis synthesize a catabolic ALS that displays a pH optimum of activity around 7.0 (86). Thus, the pH optimum is not a consistent distinguishing feature for catabolic versus biosynthetic ALS.

ALDC.

ALDC (EC 4.1.1.5) converts acetolactate to D-(−)-acetoin plus carbon dioxide. It has been purified from E. aerogenes (135), Lactobacillus casei (178), Brevibacterium acetylicum (168), and Bacillus brevis (217). As for ALS, the pH optimum of the Enterobacter ALDC is at slightly acidic pH, between pH 6.2 and 6.4. ALDC, like ALS, is functional as a dimer (135). The enzyme appears to be restricted to prokaryotes (68).

AR.

AR (EC 1.1.1.5) catalyzes both the NADH-mediated reduction of acetoin to 2,3-butanediol and the reduction of diacetyl to acetoin (Fig. 9). It is also known as butanediol dehydrogenase or diacetyl reductase and belongs to the family of zinc-independent short-chain AdhEs. The enzyme has been purified from E. aerogenes and is a homotetramer of 25-kDa subunits (210). Conversion of acetoin to butanediol is reversible. Consequently, when the NAD⁺/NADH ratio is high, AR acts as a butanediol dehydrogenase. Curiously, the reduction of diacetyl to acetoin by AR appears to be largely irreversible. As with most NADH-linked dehydrogenases, the pH optimum for activity depends on the direction of the reaction. It lies between pH 4 and 7 with diacetyl or acetoin as a substrate and around pH 9.5 with 2,3-butanediol as a substrate (210).

There are three stereoisomeric forms of 2,3-butanediol: the meso, D-(−), and L-(+) forms. They are all produced at different levels by different microorganisms. Klebsiella pneumoniae has been shown to produce between 5 and 14% of the L-(+) enantiomer, the remainder being meso-2,3-butanediol (reviewed by Hohn-Bentz and Radler [84]). Different schemes have been proposed to account for the formation of these stereoisomers (84). The situation in Klebsiella may involve an L-(+)-specific AR isoenzyme responsible for forming the minor amount of L-(+)-2,3-butanediol. A racemase must be postulated to account for the interconversion of the D and L isomers of acetoin, since ALDC generates exclusively the D-(−) isomer of acetoin (84). Indirect evidence for an L-(+)-specific AR in Klebsiella is the finding that the organism cannot oxidize D-(–)-butanediol (84).

Regulation of 2,3-Butanediol Formation.

The genes coding for the three enzymes of the 2,3-butanediol pathway in Klebsiella terrigena and E. aerogenes have been characterized (17). They form an operon (budABC) in which budA encodes ALDC, budB encodes ALS, and budC encodes AR. The presence of a diacetyl (acetoin) reductase in E. coli has also been reported (203). However, this activity was dependent on NADPH, not on NADH, which serves as an electron donor for the true ARs of the butanediol system, and no physiological role was ever proposed. It seems likely in retrospect that a side activity of another dehydrogenase may have caused the effect, perhaps of another member of the widely distributed family of short-chain Zn-independent AdhEs. A BLASTP search run by the authors failed to find any proteins closely related to BudA, BudB, or BudC on the chromosomes of E. coli or Salmonella spp.

2,3-Butanediol formation in enteric bacteria, such as Enterobacter and Klebsiella spp., requires anaerobiosis, an acidic pH, and the presence of acetate in the culture medium (18, 94, 208, 211). Acetate appears to play the key role in the induction of the budABC operon coding for the enzymes of the butanediol pathway. It is likely that anaerobiosis and low pH exert their effects via acetate accumulation in the cytosol (149). Thus, anaerobiosis should increase the rate of acetate formation from glucose, and in addition, the influx of acetate into the cells would be favored by low pH. The regulatory effect of acetate is mediated via the BudR regulatory protein, which belongs to the LysR family of regulators (149). The budR gene is separated from the budABC operon by an untranslated region of 106 bp and is transcribed in the opposite direction.

The regulation of the butanediol pathway in B. subtilis resembles that of the enterobacteria in being induced by acetate (86, 181). The alsSD operon, in which alsS encodes ALS and alsD encodes ALDC, is also under the control of the AlsR regulatory gene. Inactivation of alsR leads to the loss of expression of the alsSD operon. In addition, acetate appears to directly affect the activity of ALS. Acetate stimulates the activity of ALS but inhibits oxidation of 2,3-butanediol to acetoin at the enzyme activity level (211).

When E. aerogenes grows anaerobically in batch culture on glucose, acetate is formed via the PTA/ACK pathway, regenerating ATP. This causes acidification of the medium and cessation of growth as the pH drops below 6. The butanediol pathway is induced by the accumulated acetate, which results in formation of acetoin and 2,3-butanediol until the glucose is exhausted and the pH rises to ~6.5. During this period, >50% of the pyruvate is channeled into the formation of these neutral end products with concomitant NAD⁺ regeneration (94). Some reoxidation of 2,3-butanediol to acetoin may occur later; thus, 2,3-butanediol can serve as an external reservoir of reducing equivalents of redox potential equivalent to that of NADH (94, 137).

ACKNOWLEDGMENTS

We are grateful to Andreas Becker and Roger Harris for help in making figures.

Work in the laboratory of R.G.S. is supported by the BBSRC. Work in the laboratory of D.P.C. is supported by the U.S. Department of Energy, Office of Basic Energy Sciences.

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