Two-Carbon Compounds and Fatty Acids as Carbon Sources
DAVID P. CLARK1* AND JOHN E. CRONAN2
[SECTION EDITOR: AUGUST BÖCK]
Posted October 7, 2005
Department of Microbiology, Southern Illinois University, Carbondale, Illinois 62901,1 and Departments of Microbiology and Biochemistry, University of Illinois, B103 CLSL, 601 S. Goodwin Avenue, Urbana, Illinois 618012
*Corresponding author. Phone: (618) 453-3737, Fax: (618) 453-8036, E-mail:
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This chapter concerns the uptake and degradation of those substrates that are wholly or largely converted to acetyl coenzyme A (acetyl-CoA) in the first stage of metabolism. These include acetate, acetoacetate, butyrate, and longer fatty acids in wild-type cells plus ethanol and some longer alcohols in certain mutant strains. Entry as acetyl-CoA has two important general consequences. First, generation of energy from acetyl-CoA requires operation of both the citric acid cycle and the respiratory chain to oxidize the NADH produced. Hence, acetyl-CoA serves as an energy source only during aerobic growth or during anaerobic respiration with such alternative electron acceptors as nitrate or trimethylamine oxide. In the absence of a suitable oxidant, acetyl-CoA is converted to a mixture of acetic acid and ethanol by the pathways of anaerobic fermentation. Catabolism of acetyl-CoA via the citric acid cycle releases both carbon atoms of the acetyl moiety as carbon dioxide and growth on these substrates as sole carbon source therefore requires the operation of the glyoxylate bypass to generate cell material (Fig. 1).
The pair of related two-carbon compounds, glycolate and glyoxylate, are also discussed. However, despite having two carbons, these are metabolized via malate and glycerate, not via acetyl-CoA. In addition, mutants of Escherichia coli capable of growth on ethylene glycol metabolize it via the glycolate pathway rather than via acetyl-CoA. We have also discussed propionate as its metabolism is in many respects analogous to that of acetate.
E. coli and Salmonella enterica regard all of these two-carbon compounds as substrates of low status, and most other carbon sources (sugars, sugar derivatives, glycerol, lactate, citric acid cycle acids, etc.) are preferred growth substrates. Consequently, the expression of the genes for two-carbon substrate utilization is usually subject to strong catabolite repression.
Most of the substrates discussed in this chapter enter central metabolism as acetyl-CoA. In the simplest case, growth of wild-type Escherichia coli on acetate as the carbon source requires either acetyl-CoA synthetase or the sequential action of acetate kinase and phosphotransacetylase to convert acetate to acetyl-CoA (20, 100). Growth on the more complex substrates, acetoacetate and fatty acids, occurs by cleavage of these molecules to acetyl-CoA. Growth on the reduced substrate ethanol by adhC mutants of E. coli (which are constitutive for alcohol dehydrogenase) proceeds by oxidation of ethanol to acetyl-CoA (32, 33). In all cases the acetyl-CoA is metabolized via the citric acid cycle. The glyoxylate bypass (Fig. 1) provides the means to replenish the citric acid cycle with those intermediates lost to amino acid and heme biosynthesis (54, 93). The net effect of the glyoxylate bypass is the formation of one molecule of dicarboxylic acid from two molecules of acetyl-CoA. The glyoxylate bypass is the only anaplerotic pathway allowing growth on acetyl-CoA and is present in all organisms that utilize fatty acids or acetate as sole carbon source (93). The glyoxylate bypass will be described in detail in Chapter (Tricarboxylic Acid Cycle and Glyoxylate Bypass), but an overview is provided here.
The two unique enzymes of the glyoxylate bypass (Fig. 1), isocitrate lyase and malate synthase A, are normally induced when E. coli is grown on acetate or fatty acids (93, 176). The structural genes for isocitrate lyase (aceA) and malate synthase A (aceB) map at min 91 on the E. coli K-12 chromosome and are transcribed together with a third gene, aceK (Table 1). An adjacent gene, iclR, encodes a repressor that regulates aceBAK transcription, and iclR null mutants produce constitutive levels of isocitrate lyase and malate synthase A. The fadR gene (see below) also regulates aceBAK transcription. Transcriptional regulation is only one aspect of the regulation of the glyoxylate bypass. A major regulatory role is played by posttranslational modification of isocitrate dehydrogenase (ICDH) catalyzed by the bifunctional kinase/phosphatase encoded by the aceK gene. Addition of a phosphate to ICDH decreases its activity, thus allowing isocitrate lyase to compete more effectively for the mutual substrate, isocitrate. As will be detailed in Chapter (Tricarboxylic Acid Cycle and Glyoxylate Bypass), the function of AceK is thus to partition the flow of carbon between the glyoxylate bypass and the citric acid cycle.
Table 1Genes of glycolate and glyoxylate metabolism |
We will consider both growth on acetate as the sole carbon and energy source and incorporation of acetate when provided as a supplement for growth (Fig. 2). (Mutants of E. coli deficient in pyruvate dehydrogenase require supplementation with acetate during aerobic [but not anaerobic] growth, and mutants deficient in pyruvate formate lyase require supplementation with acetate during anaerobic [but not aerobic] growth.)
The utilization of acetate, whether for oxidation via the citric acid cycle Chapter (Tricarboxylic Acid Cycle and Glyoxylate Bypass), for replenishing intermediates of the citric acid cycle via the glyoxylate shunt Chapter (Tricarboxylic Acid Cycle and Glyoxylate Bypass), or for lipid Chapter (Biosynthesis of Membrane Lipids) or leucine Chapter (Biosynthesis and Regulation of the Branched-Chain Amino Acids) biosynthesis, requires that acetate first be activated to acetyl-CoA. Two routes are known that bring about this conversion. The first is the successive operation of acetate kinase and phosphotransacetylase and the second is acetyl-CoA synthetase.
The mechanism for the uptake and/or excretion of acetate is poorly understood. Originally, two separate transport systems were proposed, based on kinetic evidence (65). More recently, the yjcG gene, which is cotranscribed with acs, has been shown to encode an acetate uptake carrier and has been renamed actP (60). These authors also propose the existence of a second transport system for excretion and uptake of acetate.
In the acetate kinase-phosphotransacetylase pathway (Fig. 2) acetate is first converted to acetyl phosphate with cleavage of ATP to ADP and the acetyl moiety is then transferred from acetyl phosphate to CoA, with liberation of inorganic phosphate. Acetate kinase (ATP acetate phosphotransferase; EC 2.7.2.1) encoded by the ackA gene, catalyzes the first reaction and phosphotransacetylase (acetyl-CoA:orthophosphate acetyltransferase; EC 2.3.1.8) encoded by the pta gene, the second reaction. These reactions are also involved in the production of acetate during fermentation. The ackA-pta pathway is used both during anaerobic growth on sugars when acetate is a major fermentation product and during aerobic growth on acetate as carbon source (20, 98). In consequence, the biochemistry, genetics, and regulation of acetate kinase (AckA) and phosphotransacetylase (Pta) are discussed in detail in Chapter (Fermentative Pyruvate and Acetyl-Coenzyme A Metabolism). The levels of acetate kinase and phosphotransacetylase in extracts of wild-type E. coli and S. enterica vary little with different carbon sources (20, 98), and expression of the ackA and pta genes is thus neither induced by acetate nor catabolite repressed by glucose. The levels of both enzyme activities are similar (they vary less than twofold) in aerobic and anaerobic cultures (20, 98).
Under aerobic growth conditions mutants defective in either the ackA or the pta gene are severely impaired in the utilization of acetate as sole carbon source, and when grown on glucose, these strains fail to incorporate labeled acetate (20). However, ackA and pta mutants grown on glycerol are capable of incorporating labeled acetate (20). These early results indicated the presence of a second acetate incorporation system present in glycerol-grown cells but missing in glucose-grown cells. Brown et al. (20) showed that E. coli contains an inducible acetyl-CoA synthetase (Acs) that enables ackA and pta mutants to incorporate labeled acetate. Acetyl-AMP (also called acetyl-adenylate) is formed as a fairly stable intermediate that is largely enzyme bound. Acetyl-CoA synthetase (acetate:CoA ligase [AMP forming]; EC 6.2.1.1) catalyzes two successive reactions (Fig. 2). Acetate and ATP react to form acetyl-AMP, which is then attacked by the thiol of CoA to give acetyl-CoA. The lack of acetate incorporation and acetyl-CoA synthetase activity in cells grown on glucose (20) suggested that expression of this enzyme is regulated by catabolite repression and subsequent analyses have confirmed this suggestion (see below). In addition, Acs is not normally expressed anaerobically. The possession of two independent pathways means that pta/ackA mutants of E. coli and S. enterica still grow aerobically on acetate, albeit more slowly. This second route appears to be less active in S. enterica, since ackA mutants of this organism show much less residual growth on acetate (100). However, certain other enteric bacteria, such as Enterobacter aerogenes, apparently lack Acs and rely solely on the Pta/AckA pathway for aerobic growth on acetate (20, 21).
Strains of E. coli having disruptions of the chromosomal acs gene grow poorly on acetate as sole carbon source despite having a functional ackA-pta system (97). Mutants with lesions in all three genes (acs, ackA, and pta) are completely unable to grow on acetate as sole carbon source (97). This result indicates that acetyl-CoA synthetase plays a more central role in acetate metabolism than previously supposed and that no third pathway of acetate assimilation is present. It should also be noted that the acs-encoded acetyl-CoA synthetase is active with propionate and is most likely the enzyme that allows this compound to enter metabolism in both E. coli and S. enterica (19, 167)
Acetyl-CoA synthetase (Acs) is a high-affinity enzyme that allows cells to scavenge for acetate during carbon starvation and/or in the absence of a favored carbon source such as glucose. In particular, acetate excreted during rapid exponential growth on sugars may subsequently be taken back in via Acs (95, 96, 152). Cells induce the acs gene in response to rising cyclic AMP levels and falling oxygen partial pressure, as mediated by the Fnr regulator (95). Expression of the acs gene is activated by cyclic AMP and its receptor protein, Crp (9). The acs gene has two promoters, P1 (distal) and P2 (proximal) (9). Transcription of the acs gene is activated by binding of the Crp protein to tandem sites upstream of the major P2 promoter, centered at positions –69.5 and –122.5 (9, 22). Both Fis and integration host factor (IHF) also bind to the acs promoter region and reduce expression (22, 23). Fis competes with Crp for binding to overlapping sites whereas IHF binds independently. It is suggested that IHF and Fis act at different stages of growth to modulate the extent of transcription (22). The acs gene is adjacent to nrfA (nitrite reductase) and transcribed divergently. Although their regulatory regions overlap, the two genes are nonetheless regulated independently in response to quite distinct signals (23).
The flux of carbon through acetate-associated pathways is also important in acs expression. Expression is affected, probably indirectly, by the glyoxylate regulator, IclR, and its activator, FadR (96, 152). Despite being induced during stationary phase and in response to spent medium (152), transcription of the acs gene is not directly dependent on the stationary phase sigma factor, RpoS (95). Nonetheless, acs expression is somewhat reduced in rpoS mutants (152) presumably via an indirect effect. The expression of acs (together with the glyoxylate bypass) is also positively regulated by the CsrAB regulatory system (180). Mutants defective in both CsrA and RpoS grow very poorly on acetate and are also inhibited by addition of acetate to rich media (180).
Acetyl-CoA synthetase activity is regulated both at the transcriptional level, largely in response to carbon source via the Crp/cyclic AMP system, and also at the level of enzyme action by covalent modification (acetylation/deacetylation) (164, 166). Sirtuins are NAD-dependent protein deacetylases that were originally discovered in higher organisms where they are involved in deacetylation of histones that are acetylated on lysine residues. The NAD is cleaved and the acetyl group removed from the protein substrate is attached, generating the novel metabolite O-acetyl-ADP-ribose (150). Sirtuins are found in all forms of life, including bacteria. The CobB protein is a bacterial sirtuin that acts in an analogous manner but whose substrate is acetyl-CoA synthetase. CobB activates the enzyme activity of Acs by removing an acetyl group from an active-site lysine residue (K609) (163, 165). Acetylation of K609 of Acs is due to a protein acetyltransferase called Pat, which uses acetyl-CoA as a source of acetyl groups (165). Thus Acs activity is regulated posttranslationally by reversible acetylation and deacetylation. Acetylation inactivates the enzyme by blocking synthesis of the acetyl-AMP intermediate, although it has no effect on the thioester-forming activity of the enzyme (163). The enzyme is thought to undergo a large conformational change in the transition between the two partial reactions (66). The physiological relevance of sirtuin control of Acs remains to be elucidated.
A link between acetate metabolism and chemotaxis has been known for some time. The chemotaxis protein CheY interacts with the flagellar switch so causing clockwise rotation and this activity of CheY is increased by acetylation of the protein. Originally acetyl-AMP (made by Acs) was thought to acetylate CheY (7). However, it was later found that acetyl-CoA synthetase itself directly acetylates CheY (4, 5, 6). Acs can use either acetate (plus ATP) or acetyl-CoA as a source of acetyl groups. Acs acetylates itself and then transfers the acetyl group to CheY (6). Acetylated CheY may have from one to six acetyl groups attached to different lysine residues. CheY may also be phosphorylated by the CheA histidine kinase and acetyl phosphate. Phosphorylation and acetylation of CheY appear to be mutually exclusive (5). The precise regulatory roles of these modifications are uncertain, but they are thought to link chemotaxis in some way to the metabolic state of the cell. Mutant strains of E. coli lacking Acs showed much reduced sensitivity to chemotactic stimuli (both attractants and repellents) (4).
It has been known anecdotally for some time that there are significant differences in acetate metabolism among different laboratory strains of E. coli. Recently, the differences between two commonly used lines, BL21 and JM109, were more fully investigated (52, 115, 138). Strain BL21 excretes relatively little acetate during growth on glucose, whereas strain JM109 shows the opposite. In strain BL21 the glyoxylate cycle is anomalously active during growth on glucose (52, 115). Normally the aceBAK operon is repressed by carbon sources other than acetate and the glyoxylate cycle is scarcely active. In addition, acetyl-CoA synthetase is expressed at much higher levels in strain BL21 (138). Thus strain BL21 tends to convert surplus acetyl-CoA to malate, via the glyoxylate bypass, rather than excreting it as acetate, via Ack and Pta. In addition, its higher Acs levels suggest that strain BL21 may be reincorporating acetate, even during growth on glucose. Whether such strain differences are due to single or multiple genetic differences is still uncertain. However, no transcription of the iclR gene was observed in strain BL21 (138) and it is possible that most of the metabolic perturbations are direct or indirect consequences of this. Engineered up-regulation of the glyoxylate bypass by introducing a defect in fadR and overproduction of phosphoenolpyruvate carboxylase (thereby routing pyruvate to oxaloacetate rather than acetyl-CoA) have both been shown to significantly reduce acetate excretion (52).
The emergence of DNA microarray technology has allowed the global assessment of gene regulation in response to a variety of stimuli, including environmental conditions. Although valuable, the data from this approach are still compromised by technical difficulties. Unfortunately, results from different laboratories that assayed the same organism under the same conditions often conflict (107). Several investigators have compared gene expression in E. coli grown on different carbon sources. When cells grown on acetate are compared with those grown on glucose, transcription of an estimated 300 to 400 genes is increased and approximately the same number are down-regulated (124, 125, 172). Apart from questions of reproducibility, interpretation of these data is difficult since induction by "growth on acetate" actually involves multiple factors including (i) growth using acetate as carbon source per se, (ii) increased gluconeogenesis and decreased glycolysis due to growth in the absence of sugars or their derivatives, (iii) increased activity of the respiratory chain and citric acid cycle (common to growth on nonfermentable carbon sources such as acetate or citric acid cycle intermediates), (iv) decreased catabolite repression (common to growth on a variety of poor carbon sources), and (v) growth rate regulation due to growth on acetate being very slow relative to growth on most other carbon sources, especially sugars. Another major problem is acetate toxicity. Many genes induced by acetate are shared with the responses to other weak permeant acids such as propionate and benzoate (139). Finally, there are a series of poorly understood responses to acetate. For example, operation of the PhoPQ two-component system, which responds to divalent cations and pH, is inhibited by acetate. Apparently the buildup of acetyl-CoA inhibits autophosphorylation of the PhoQ protein (99).
Here, we are concerned primarily with the catabolic pathways required for utilization of acetate; consideration of the other factors is beyond the scope of this chapter. Overall, growth on acetate results in the following increases in gene expression that are clearly related to acetate metabolism per se (124, 125, 172). Large (fivefold or greater) increases are seen in expression of genes encoding the glyoxylate bypass enzymes (aceBAK), acetyl-CoA synthetase (acs), phosphoenolpyruvate synthase (ppsA), pyruvate carboxykinase (pckA), and aldehyde dehydrogenase (aldA). Moderate (two- to fivefold) increases in expression are seen in genes encoding several citric acid cycle enzymes, fructose biphosphate synthase (fbp), and the NADP-linked malic enzyme (maeB).
Growth on acetate also results in moderately decreased expression (two- to fivefold) for the genes encoding acetate kinase and phosphotransacetylase (ackA and pta). Moreover, expression of many genes encoding proteins involved in glycolysis and the transport and metabolism of sugars is similarly decreased. Many genes for biosynthetic enzymes also show moderately decreased expression, presumably in response to the slow growth rate on acetate. Three enzymes, phosphoenolpyruvate synthase, pyruvate carboxykinase, and fructose biphosphate synthase, are needed for gluconeogenesis during growth on acetate. The NADP-linked malic enzyme is presumably needed for pyruvate production. The twofold down-regulation of the ackA and pta genes seen during growth on acetate is consistent with other data showing that acetate is largely catalyzed by acetyl-CoA synthetase under these conditions.
During growth on acetate excess NADPH is generated (largely by the NADP-linked isocitrate dehydrogenase of the citric acid cycle and the NADP-linked malic enzyme). Indeed, the surplus must be removed to regenerate NADP+. This is done by the soluble transhydrogenase, UdhA, which uses the surplus NADPH to convert NAD+ to NADH (149). UdhA is essential for growth on acetate (and under other conditions resulting in NADPH overproduction) (149). (The better- known transhydrogenase, PntAB, which is membrane bound and energy dependent, is used in the converse situation, to generate NADPH using NADH.) Expression levels of pntAB and udhA vary in response to NADPH levels. The level of UdhA rises when the NADPH/NADP+ ratio is high and falls when it is low, whereas the level of PntAB changes in the opposite direction (149). The citric acid cycle of eukaryotic mitochondria has an isocitrate dehydrogenase that uses NAD, whereas most bacteria contain an NADP-linked enzyme (189). This NADP-linked isocitrate dehydrogenase of the citric acid cycle is necessary to generate NADPH during growth on acetate, presumably because of the relatively low flux through the NADPH-generating steps glucose-6-phosphate dehydrogenase and gluconate-6-phosphate dehydrogenase that lead into the pentose cycle. When the isocitrate dehydrogenase of E. coli was engineered to use NAD instead of NADP, growth on acetate was greatly impaired (189). A survey of prokaryotic genomes shows that organisms with genes for the glyoxylate bypass always possess NADP-linked isocitrate dehydrogenase, whereas organisms having NAD-linked isocitrate dehydrogenase never possess isocitrate lyase (189).
The following cautionary tale shows how genes needed under particular growth conditions may have roles that are not immediately obvious. This caution is especially relevant in these days of microarray analyses. Mutants of E. coli defective in the ilvBN genes that encode acetohydroxy acid synthase I (AHAS-I) can still synthesize the amino acids isoleucine and valine because of the presence of a second isoenzyme, AHAS III (encoded by ilvIH). Consequently, these strains can grow on glucose or succinate minimal medium without addition of isoleucine plus valine. However, ilvBN mutants fail to grow on acetate or fatty acids as carbon source unless supplemented with isoleucine plus valine (42, 43). The reason for this phenomenon is that the intracellular concentration of pyruvate, an essential AHAS substrate, is low during growth on acetate and AHAS-I is the only AHAS that does not strongly favor α-ketobutryate over pyruvate (8, 42, 43, 62). A similar situation is seen in S. enterica, but for different reasons. Both E. coli K-12 and S. enterica LT2 have AHAS-I. However, the second AHAS isozymes differ in the two organisms; E. coli has AHAS-III whereas S. enterica has AHAS-II. In E. coli the effect of acetate on branched chain amino acid synthesis has been attributed to the preference of AHAS-III for α-ketobutryate over pyruvate, whereas the inability of AHAS-II to function when acetate is the carbon source is due to its preference for α-ketobutryate plus inhibition of the enzyme by the glyoxylate that accumulates during growth on acetate (51).
Several other roles, in addition to being an intermediate in the conversion of acetate to acetyl-CoA by the AckA/Pta pathway, have been proposed for acetyl phosphate. The possible regulatory effects of acetyl phosphate are discussed in Chapter (Tricarboxylic Acid Cycle and Glyoxylate Bypass). Acetyl phosphate was once proposed as the energy source for those active transport systems using periplasmic binding proteins (73). However, these observations were not confirmed, and strains carrying deletions of the pta-ack operon are not noticeably deficient in the uptake of nutrients accumulated by these systems. The observation that a phosphoryl group may be transferred from acetate kinase to enzyme I of the phosphotransferase system may be relevant (53). This route for phosphoryl transfer implies that acetyl phosphate can donate its high-energy phosphate to the phosphotransferase system via acetate kinase and enzyme I and hence provide an alternative energy source to phosphoenolpyruvate (53). Such a situation seems likely to occur only during anaerobic conditions when acetyl phosphate is being generated rapidly by sugar fermentation Chapter (Fermentative Pyruvate and Acetyl-Coenzyme A Metabolism).
The arpA (previously yjaC) gene is located between the aceBAK operon and the iclR regulatory gene. Sequence analysis indicates the presence of ankyrin repeats in the ArpA protein. When human RNase L was expressed in E. coli, it prevented growth on acetate, and the enzymes of the glyoxylate bypass were not induced (44). The fact that RNase L and ArpA both have ankyrin repeats led the authors to suggest an interaction between these two proteins and to postulate that ArpA was somehow needed for induction (44). However, an interaction between RNase L and another ankyrin repeat protein, YahD, has also been suggested as the mechanism of RNase L toxicity (131). However, there is no direct evidence for either interaction. There is also, as yet, no direct evidence (despite its provocative chromosomal location) that ArpA plays a role in acetate metabolism. Considerable sequence-length polymorphism exists in the aceBAK-iclR region, and the presence of the arpA gene, located between aceK and iclR, varies greatly among natural isolates of E. coli (113). Curiously, the absence of arpA seems to correlate at least to some extent with virulence (36). It has been suggested that, while not a virulence determinant as such, an operational glyoxylate cycle is necessary for survival of many pathogenic microorganisms within the host (103).
Enteric bacteria such as E. coli and S. enterica produce ethanol during fermentation by reducing the acetyl group of acetyl-CoA to acetaldehyde and then to ethanol (Fig. 2). Both reactions consume NADH. The first reaction is catalyzed by CoA-linked acetaldehyde dehydrogenase (ACDH) and the second is catalyzed by alcohol dehydrogenase (ADH). A single large protein, AdhE, is responsible for both activities (40, 63). This polypeptide of 891 amino acids has an N-terminal domain responsible for the ACDH activity and a C-terminal domain where the ADH activity is located. Sequence comparisons suggest that the adhE gene of E. coli may be the product of the fusion of two genes, one encoding ADH and the other encoding ACDH (33). (Further details of AdhE and alcohol production are discussed in Chapter (Fermentative Pyruvate and Acetyl-Coenzyme A Metabolism).) When ethanol is used as a carbon source by aerobic organisms, the two reactions above occur in reverse. The resulting acetyl-CoA is oxidized via the citric acid cycle, and the NADH produced is oxidized by the oxygen-linked respiratory chain.
Although E. coli is a facultative anaerobe and can grow in air by oxidizing such substrates as acetate, succinate, or glycerol, wild-type strains are unable to grow aerobically at the expense of ethanol or other short-chain alcohols. The reasons for this are twofold. First, the adhE gene encoding the bifunctional AdhE protein with both ADH and ACDH activities is poorly expressed under aerobic conditions (32). Second, AdhE activity is highly oxygen sensitive and is inactivated in the presence of air. Nonetheless, mutants (designated "adhC") were isolated that were capable of growing aerobically on ethanol or 1-propanol as carbon source (35). These strains proved to contain two closely linked mutations, one within the promoter region and a second in the structural gene for adhE (see Chapter Fermentative Pyruvate and Acetyl-Coenzyme A Metabolism). Alterations in the promoter region resulted in the expression of high levels of AdhE under aerobic conditions. These mutations proved to be in the binding site for the Cra regulator, which normally represses adhE expression. In the adhC mutants, Cra no longer bound and adhE was consequently derepressed (83).
The mutations in the structural adhE gene resulted in an altered protein that was no longer oxygen sensitive. This aerotolerant form of AdhE has two amino acid residue changes, A267T and E568K. However, the E568K substitution is sufficient to protect the enzyme against oxidation. Furthermore, replacement of E568 by virtually any other residue resulted in AdhE isoforms that were active under both aerobic and anaerobic conditions (72, 108). 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 (A267T, E568K) against metal-catalyzed oxidation (50).
The AdhE multifunctional enzyme is highly active with both two- and three-carbon substrates and will therefore convert ethanol to acetyl-CoA and 1-propanol to propionyl-CoA. Consequently, adhC mutants will also grow aerobically on 1-propanol as sole carbon source. Longer-chain substrates are less well metabolized, although there is a significant difference between the enzymes from E. coli and S. enterica. The S. enterica AdhE works reasonably well with substrates up to 6 to 8 carbons in length (see Chapter Fermentative Pyruvate and Acetyl-Coenzyme A Metabolism) for details). The E. coli enzyme shows moderate activity against 1-butanol, but activity is extremely poor with substrates of five carbons or longer. Although they are able to convert butanol to butyryl-CoA via AdhE, adhC mutants of E. coli fail to grow on 1-butanol as sole carbon source. One reason is the low activity of AdhE with butanol as substrate. A second problem is that wild-type strains of E. coli cannot metabolize butyryl-CoA or butyrate. However, double mutants able to grow on butyrate are known. Such mutants have lesions in both the fadR and atoC regulatory genes and hence express both the acetoacetate pathway and the β-oxidation system for fatty acids in a constitutive manner (see below).
In such strains, butyrate is converted to butyryl-CoA by the atoAD-encoded transferase (Fig. 3). The oxidation of butyryl-CoA to crotonyl-CoA by FadE, the hydration of crotonyl-CoA to acetoacetyl-CoA by FadB, and finally the AtoB-catalyzed thiolytic cleavage of acetoacetyl-CoA yields two molecules of acetyl-CoA. The requirement for mutations in both atoC and fadR is due to the inability of butyrate to induce either the ato operon or the fad regulon. Since an atoC fadR double mutant can use butyrate, it might be supposed that an atoC fadR adhC strain would grow on butanol. However, this is not the case, apparently because the activity of AdhE is insufficient to effectively convert butanol to butyryl-CoA (35). However, starting from adhC strains, mutants that possess 10-fold further elevated levels of AdhE can be isolated. Such strains (e.g., DC430) were originally selected by growth on ethanol in the presence of the ADH inhibitor 4-methyl-pyrazole (35). Such mutants possess much higher levels of adhE messenger RNA because of a defect in ribonuclease G. It turns out that RNase G is involved not only in rRNA processing but also in the turnover of the mRNAs of a few metabolic genes, in particular, adhE and eno (encoding enolase). The rng430 mutation of DC430 has a single amino acid alteration, G341S, which results in an altered RNase G that retains the ability to process 16S rRNA but no longer degrades adhE mRNA (178). (For more details, see Chapter Fermentative Pyruvate and Acetyl-Coenzyme A Metabolism). DC430, the original strain carrying the rng430 mutation, grew poorly on acetate or ethanol in some media, a behavior apparently resulting from acetate toxicity rather than the inability to metabolize acetate. Whether this is due to side effects of the rng430 mutation or to other mutations remains unclear. However, to restore strong growth on ethanol or acetate a further mutation was then selected. This final E. coli strain was able to grow on butanol as carbon source (35). Both the adhC and the rng430 mutations are necessary to give sufficiently high enzyme levels of ADH and ACDH to metabolize butanol to butyryl-CoA, whereas the other mutations are necessary for the metabolism of butyryl-CoA (Fig. 3).
The compounds discussed thus far are all metabolized via acetyl-CoA and the citric acid cycle and all require the glyoxylate bypass to replace citric acid cycle intermediates. Ironically, glyoxylate itself and the related two-carbon compound glycolate are exceptions to this general rule. These two compounds are products of plant and algal metabolism and are utilized as sole sources of carbon and energy by a variety of bacteria including E. coli. However, although S. enterica grows on glycolate it cannot grow on glyoxylate for unknown reasons (67). Nonetheless, the S. enterica genome sequence shows the presence of a full set of genes for glyoxylate metabolism. Glycolate can also be produced by oxidation of glycolaldehyde, which is produced by catabolism of certain carbohydrates such as D-arabinose (141) and as a by-product from DNA repair (see below). Mutants capable of growing on ethylene glycol also metabolize it via glycolaldehyde and glycolate (see below).
The first step of the glycolate pathway is the oxidation of glycolate to glyoxylate (Fig. 4), catalyzed by glycolate oxidase. Glyoxylate is then metabolized by either of two divergent condensation reactions, one leading to the glycolytic pathway and the other to the citric acid cycle (126). In the first alternative, two molecules of glyoxylate are condensed by glyoxylate carboligase to form tartronic semialdehyde plus CO2. Tartronic semialdehyde is then reduced to glycerate, which is phosphorylated to give the glycolytic intermediate glycerate 3-phosphate (126). The second alternative involves condensation of glyoxylate with acetyl-CoA to form malate, a citric acid cycle intermediate (126, 176). This reaction is identical with that of the glyoxylate cycle (Fig. 1), although the condensation is catalyzed by malate synthase G encoded by the glcB gene rather than by the glyoxalate cycle enzyme malate synthase A encoded by the aceB gene (126, 176, 177). Malate synthases G and A are regulated by distinct mechanisms. Malate synthase G is strongly induced (1,000-fold) by glycolate (108), whereas malate synthase A is induced 20-fold by growth on acetate or fatty acids (176). Mutants lacking glyoxylate carboligase fail to grow on either glycolate or glyoxylate (30), whereas mutants lacking glycolate oxidase grow on glyoxylate, but not glycolate (126). Malate synthase G is not essential for growth on either compound as sole carbon source, since glyoxylate carboligase gives rise to 3-phosphoglycerate, a glycolytic intermediate. In contrast, glyoxylate carboligase is essential even when malate synthase G is functional, because the acetyl-CoA needed for the malate synthase G reaction is derived, ultimately, from glycolysis. The dispensable role of the citric acid cycle in the metabolism of these compounds is shown by the finding that mutants deficient in citrate synthase (gltA) grow on glycolate with the same rate and yield as a wild-type strain (126). However, during growth on the more oxidized compound glyoxylate, the presence of malate synthase G allows more rapid growth of gltA strains (126).
A cluster of genes involved in glycolate metabolism is located at min 67 of the E. coli map (Table 2). The glcDEFGBA operon is transcribed in divergent orientation to the neighboring regulatory gene, glcC (135, 137). The glcDEF genes encode subunits of glycolate oxidase, glcB encodes malate synthase G, and glcA encodes a permease (116). The role of glcG is unknown and mutations in this gene do not appear to affect growth on glycolate.
Table 2Genes of glycolate and glyoxylate metabolism |
The deduced amino acid sequence of GlcB (108) is similar to that of other known malate synthases including malate synthase A (AceB) of the glyoxylate bypass. The sequences of GlcD and GlcE are similar to those of several flavoenzymes, and GlcF is similar to certain iron-sulfur proteins (135). Earlier reports suggested that the glcB gene (encoding malate synthase G) was not cotranscribed with the neighboring genes that encoded glycolate oxidase (109). However, the isolation of point mutants that simultaneously lost both of enzyme activities suggested a common regulatory system (126). Indeed, later investigations showed that these genes constitute an operon that is under control of the GlcC activator protein (136).
The genes involved in the conversion of glyoxylate to 3-phosphoglycerate are located at min 11 to 12 of the E. coli map (41) (Table 2). The gcl gene encodes glyoxylate carboligase, the glxR gene encodes tartronic semialdehyde dehydrogenase, and the glxK gene encodes glycerate kinase. The tartronic semialdehyde dehydrogenase belongs to a family of structurally and mechanistically related NAD(H)-linked β-hydroxyacid dehydrogenases (114). The hyi gene encodes hydroxypyruvate isomerase (previously known as glyoxylate-induced protein, Gip) (2). This enzyme interconverts tartronic semialdehyde and hydroxypyruvate. Glyoxylate is a product of purine metabolism as well as being formed from glycolate. Presumably due to this connection, the gcl, glxR, glxK, and hyi genes are clustered with the genes for allantoin degradation. Allantoin is the product of purine degradation and is converted to ureidoglycolate. This may be converted to oxaluric acid (by ureidoglycolate dehydrogenase) or it may be split (by ureidoglycolate hydrolase) releasing urea and glyoxylate (41).
An additional minor source of glycolate is produced by DNA repair. Oxidative damage of DNA, by agents such as ionizing radiation or bleomycin, generates 3' termini with phosphoglycolate groups that can be removed by either exonuclease I (146) or exonuclease IX (147). The released 2-phosphoglycolate is dephosphorylated to glycolate by the enzyme 2-phosphoglycolate encoded by the gph gene (147). The phosphatase is expressed constitutively (147).
Glyoxylate carboligase is curious in belonging to a class of enzymes that require FAD, although there is no oxidation or reduction step in the reaction catalyzed (30, 37). The most numerous members of this group are the acetohydroxy acid synthases catalyzing early reactions of the branched-chain amino acid synthetic pathway (see above and Chapter Biosynthesis and Regulation of the Branched-Chain Amino Acids). These enzymes catalyze a reaction chemically analogous to that of glyoxylate carboligase, and the Gcl protein shares 30% amino acid residue identity with the AHAS isozymes (30). This group of enzymes shows significant similarity to pyruvate oxidase (PoxB of E. coli) which does use FAD for the oxidation of pyruvate to acetate. Glyoxylate carboligase contains a vestigial quinone binding site, as do the acetohydroxy acid synthases (29, 30). These data, plus new data showing a side reaction of AHAS-II involving electron transfer (170), argue that glyoxylate carboligase, like the AHAS isozymes was derived from a functional FAD-linked dehydrogenase.
Duplicate genes encoding the enzymes tartronic semialdehyde reductase (garR, previously yhaE) and glycerate kinase (garK, previously yhaD) are found in the pathway for the degradation of glucarate and galactarate (76). This pathway generates tartronic semialdehyde by splitting a six-carbon intermediate (5-keto-4-deoxy-D-glutarate), and glyoxylate is not involved as an intermediate. The genes for glucarate and galactarate catabolism are under common regulation by the cdaR gene (previously sdaR and yaeG) (110). The CdaR protein is believed to be an activator that induces its target genes in response to glycerate, glucarate, or galactarate (110). Since growth on glycolate or glyoxylate generates glycerate as an intermediate, it seems likely that the garK and garR genes are induced under these conditions together with the glxR and glxK genes. Indeed, both glycerate kinases are present in cells grown on glycolate (126). In retrospect, this probably explains why the only mutants in the glyoxylate pathway isolated by physiological means were in gcl, encoding the only unique enzyme of the pathway, glyoxylate carboligase. Both of the glycerate kinases were originally believed to generate 3-phosphoglycerate (16, 118). However the GarK isoenzyme was recently shown to generate the 2-phospho isomer (18). Whether the GlxK isoenzyme also generates 2-phosphoglycerate remains unknown. There may be further isozymes of this pathway. A putative hydroxypyruvate isomerase homolog, YgbM, is located in a cluster of genes that includes a possible β-hydroxyacid dehydrogenase (YgbJ, conceivably a third tartronic semialdehyde reductase) and an aldolase (YgbL) (88, 114)
Two proteins that act as NADPH-linked glyoxylate reductases have been found in E. coli, encoded by the ycdW and yiaE genes (117). The YiaE protein was previously identified as a 2-ketoaldonate reductase of broad specificity. YiaE converts 2,5-diketo-D-gluconate to 5-keto-D-gluconate, 2-keto-D-gluconate to D-gluconate, and 2-keto-L-gulonate to L-idonate (188). Disruption of the chromosomal yiaE gene resulted in the loss of 2-ketoaldonate reductase activity and loss of the ability to use 2-keto-D-gluconate as carbon source (188). Both YiaE and YcdW can use both glyoxylate and hydroxypyruvate; however, the YcdW is most active with glyoxylate, whereas YiaE prefers hydroxypyruvate. Since reduction of glyoxylate generates glycolate and since glycolate is, as far as it is known, always metabolized via glyoxylate, the physiological role of glyoxylate reductase is problematical. Conceivably, it might act to generate reduced NADPH under certain circumstances. On the other hand, we may not know the true physiological substrate(s) for YcdW and YiaE. The two glyoxylate reductases appear to be constitutive (117).
Glycolate is known to induce glycolate oxidase and malate synthase G (109, 126, 176), whereas only glyoxylate, the product of glycolate oxidase, induces glyoxylate carboligase (30, 126). This is the result of control of the glycolate gene cluster by GlcC (135), whereas the gcl gene is in the allantoin/glyoxylate cluster under control of AllR and AllS (41). The increased enzyme activities observed upon addition of glycolate or glyoxylate are due to increased transcription of the structural genes. The GlcC protein acts as a positive regulator and mediates induction of the glcDEFGBA operon in response to glycolate (137). Expression of the glc operon requires an IHF and is repressed by the ArcAB regulatory system. The glcC regulatory gene is expressed constitutively (137).
The allantoin/glyoxylate regulon is controlled by two regulatory genes, allR (41) and allS (previously ybbS) (143), each of which is transcribed from its own promoter. The structural genes comprise three operons: allA, gcl-hyi-glxR-allP-allB-ybbY-glxK, and (transcribed in the opposite orientation) allD-allC-ylbA. The AllR repressor controls all three structural gene operons. Deletion of the allR gene results in constitutive expression of the regulon indicative of negative regulation (41). In wild-type strains, the allantoin regulon is derepressed in the presence of allantoin or glyoxylate. However, the true inducer appears to be glyoxylate since, in mutants unable to convert allantoin to glyoxylate, only glyoxylate acts as inducer (143). Furthermore, binding of AllR to promoter fragments as monitored by DNA gel-shift experiments was abolished in the presence of glyoxylate (but not allantoin or glycolate) (143). In addition, AllR represses the allS gene. This is because the allS and allA genes are divergently transcribed from promoters that share a single operator site that binds AllR (143). The role of AllS is to activate expression of the allD operon that encodes a branch pathway for generation of ammonia from allantoin under anaerobic conditions. This pathway converts ureidoglycolate to oxaluric acid instead of glyoxylate. When the allS gene is deleted, expression of the allD operon is abolished (143). The AllS protein does not affect the other two operons involved in allantoin/glyoxylate metabolism. The allS gene itself is expressed anaerobically and is hyperinduced during nitrogen deficiency. However, nitrogen regulation is not due to NtrC or Nac. The AllS protein binds to the allD promoter in the presence or absence of allantoin (143).
Glyoxylate is a common intermediate of the glycolate pathway (encoded by glcDEFGBA) and the glyoxylate cycle (encoded by aceBAK). The two operons are cross-induced. That is, the presence of either acetate or glycolate induces both operons (137). In the case of the glc operon cross-induction is mediated by the GlcC protein, which recognizes acetate as an alternative effector (137). Both operons are also affected similarly by IHF and by ArcAB. One consequence of this cross-induction is that both malate synthase isoenzymes (GlcB or AceA) are expressed simultaneously in cells grown on either glycolate or acetate. Hence, null mutations in either glcB or aceA alone show no growth phenotype (137). It seems that glyoxylate produced during operation of the glyoxylate shunt can also induce gcl expression since increased glyoxylate carboligase activity is seen during growth on acetate (126).
Three alternative carriers, GlcA, LldP, and ActP, are thought to mediate transport of glycolate (60, 116). The GlcA protein (previously YghK) is the "natural carrier" in the sense that it is regulated in response to the presence of glycolate and encoded by the glcA gene, which is located in the glc operon (118). LldP protein (L-lactate permease) belongs to the L-lactate regulon and ActP (previously YjcG) is encoded in the same operon as Acs and presumably intended for acetate transport (60). Both GlcA and the LldP are equally capable of high-affinity transport of glycolate, L-lactate and D-lactate. Mutual competition between these substrates showed Ki values for all three in the 10 to 20 μM range. Both transport systems are driven by the proton motive force as indicated by sensitivity to inhibition by the uncoupler, carbonyl cyanide m-chlorophenylhydrazone (CCCP) (116). Sequence comparisons indicate that ActP is a member of the sodium:solute symporter family (60).
Wild-type E. coli cannot use ethylene glycol as a carbon source, and mutant strains able to use this glycol could not be directly selected in a wild-type strain. However, such strains were obtained starting from a mutant strain able to use 1,2-propanediol (i.e., propylene glycol) (17). Two major regulatory alterations were involved. First, levels of propanediol oxidoreductase (encoded by fucO), needed to metabolize 1,2-propanediol in the parental strain, were further elevated. This enzyme also converts ethylene glycol into glycolaldehyde. Second, constitutive high-level synthesis of an NAD-linked "glycolaldehyde dehydrogenase" that converts glycolaldehyde to glycolate was needed (17). The glycolate was then metabolized by the glycolate pathway, as described above. The "glycolaldehyde dehydrogenase" was shown to be identical with lactaldehyde dehydrogenase (EC 1.2.1.22), an enzyme involved in the aerobic metabolism of fucose (24). This enzyme is induced under aerobic conditions in wild-type cells by the deoxysugars fucose and rhamnose, whose aerobic metabolism yields lactaldehyde. It is also induced by glutamate and related metabolites. Lactaldehyde dehydrogenase is encoded by the aldA gene which is located far away from the other genes involved in fucose or rhamnose metabolism (31). Lactaldehyde dehydrogenase catalyzes the oxidation of several α-hydroxyaldehydes such as lactaldehyde, glyceraldehyde, or glycolaldehyde with a Km in the micromolar range (3)
The two-carbon dicarboxylic acid, oxalate, may be used as sole carbon and energy source by a few rare anaerobic microorganisms, such as Oxalobacter formigenes (1). This pathway involves three steps. Oxalate is taken up by an oxalate/formate antiporter, followed by transfer of the CoA moiety of formyl-CoA to one of the oxalate carboxyl groups to give oxalyl-CoA that is then decarboxylated to formyl-CoA. Energy, as proton motive force, is derived from the consumption of a proton during the transferase reaction followed by the electrogenic antiport of oxalate for formate by the oxalate-formate antiporter, OxlT (168). Although E. coli cannot grow on oxalate, it nonetheless possesses homologues of the genes involved in oxalate metabolism and therefore might participate in oxalate degradation in the intestine. The YfdW protein appears to be an oxalate:formyl-CoA transferase and YfdU may be an oxalyl-CoA decarboxylase (8). Perhaps the most interesting aspect of the pathway is the structure of the oxalate formyl-CoA transferase. This forms a dimer of two intercatenated rings with the shared CoA binding site located at the interface between the two subunits (64).
Propionate is a common carbon source present in many environments and may be utilized by many bacteria including E. coli and S. enterica. In addition, propionate, in the form of propionyl-CoA, is generated internally as an end product of the degradation of fatty acids having an odd number of carbon atoms. Propionate metabolism has the dubious distinction of being perhaps the most thoroughly confused metabolic pathway of E. coli. Several pathways for propionate metabolism are known in various bacteria (169). At one time or another, at least five pathways have been proposed for E. coli. Only recently was the methylcitrate cycle (Fig. 5) established as the route of propionate degradation, first in S. enterica (75, 171) and then in E. coli (59, 102, 169). Although this pathway has been known for some time, it was regarded as characteristic of fungi where it was first established in the 1970s.
In retrospect, a major portion of the confusion was that most of the enzymes involved in propionate metabolism also function, at least to some extent, with the corresponding substrates of acetate metabolism. The simplest example of this is that acetyl-CoA synthetase and propionyl-CoA synthetase can use both acetate and propionate, although each enzyme is most active with its eponymous substrate. Similarly, the enzymes that use methylcitrate or methylisocitrate also have some activity on citrate or isocitrate, respectively (the converse is also true in some cases).
The situation was more complicated in E. coli than in S. enterica as many strains of E. coli require the glyoxylate cycle to grow on propionate whereas S. enterica does not. The glyoxylate cycle is needed to regenerate oxaloacetate (via malate), not to metabolize propionate per se. This introduced more confusion because the glyoxylate enzymes can also function to some extent with the larger C5 homologues.
The genes for propionate metabolism form (Table 3) an operon under control of the PrpR activator, which is transcribed divergently from the clustered structural genes, prpBCDE (75, 171). Upon entering the cell, propionate is first activated by the propionyl-CoA synthetase (encoded by prpE). The propionyl-CoA then reacts with oxaloacetate to give 2-methylcitrate in a reaction analogous to the condensation of acetyl-CoA with oxaloacetate to give citrate in the citric acid cycle. This is catalyzed by 2-methylcitrate synthase (PrpC).
Table 3Genes of propionate metabolism |
The 2-methylcitrate is then isomerized to 2-methylisocitrate via 2-methyl-cis-aconitate. However, this step differs from the situation in the citric acid cycle since it requires two enzymes (87, 89, 90). The prpD-encoded 2-methylcitrate dehydratase converts 2-methylcitrate to 2-methyl-cis-aconitate. The product is rehydrated by either of the aconitase isoenzymes (AcnA and AcnB) of the citric acid cycle, which both work with either cis-aconitate or 2-methyl-cis-aconitate. The PrpD enzyme was originally misidentified as a third aconitase ("AcnC"). However, although it can dehydrate both citrate and methylcitrate, it is unable to catalyze the second stage of the aconitase reaction (15, 18, 19). PrpD shows no sequence similarity to AcnA and AcnB or other related (de)hydratases. Finally, the 2-methylisocitrate is split by 2-methylisocitrate lyase, PrpB, to give succinate and pyruvate in a reaction analogous to the isocitrate lyase reaction of the glyoxylate bypass (19, 75, 169). Recent crystal structures provide a firm structural basis for the ability of the enzyme to discriminate between isocitrate and 2-methylisocitrate (101).
Oxaloacetate is required for priming the methylcitrate cycle. In E. coli oxaloacetate is provided by the glyoxylate bypass, which is therefore required for growth on propionate, whereas in S. enterica oxaloacetate is made from pyruvate via Pps (PEP synthetase) and Ppc (PEP carboxylase). Consequently, mutants of S. enterica defective in glyoxylate cycle function grow on propionate (75), whereas mutants lacking Pps fail to grow on propionate. The Cra regulator is required for expression of both pps and ppc, and hence cra mutants of S. enterica also fail to grow on propionate (75).
The propionate operon is also required when propionyl-CoA is generated by other metabolic pathways. These include the β-oxidation of fatty acids having an odd number of carbon atoms. After pairwise removal of the carboxyl-proximal carbon atoms the fatty acid chain, the final three carbon atoms remain as propionyl-CoA. Growth by adhC mutants on n-propanol (or by adhC atoC mutants on n-pentanol; see above) will also generate propionyl-CoA. S. enterica, but not E. coli, is also capable of growth on 1,2-propanediol as a carbon and energy source. This leads to the formation of propionate, which is then converted to propionyl-CoA by one of three alternatives: (i) the propionate kinase of the 1,2-propanediol degradation system (PduW) followed by phosphotransacetylase (Pta), (ii) propionyl-CoA synthetase (PrpE), or (iii) acetyl-CoA synthetase (130).
The prpBCDE operon is controlled by the transcriptional activator PrpR. This is encoded by the prpR (previously yahP) gene which is transcribed divergently from the prpBCDE operon. Mutants lacking PrpR fail to grow on propionate, which is indicative of positive regulation (78). The nitrogen sigma factor, RpoN (also called NtrA and sigma-54), is required for expression of the prpBCDE operon and there is a consensus sigma-54 binding site in the promoter region (78). The IHF protein is also necessary for expression (130). Deletion studies on the prpR gene have shown that the N-terminal domain of PrpR is the coactivator-sensing region of the protein. A PrpR protein lacking the N-terminal domain activates the prpBCDE operon constitutively (129, 130). The true inducer of the methylcitrate cycle is 2-methylcitrate rather than propionate. The PrpR activator protein binds 2-methylcitrate but not citrate or propionate (129). Induction of the prpBCDE operon requires the initial synthesis of propionyl-CoA followed by the synthesis of 2-methylcitrate from propionyl-CoA and oxaloacetate by PrpC (171). In the absence of functional PrpC protein the prpBCDE operon becomes noninducible (171).
The source of propionyl-CoA for induction of the prpBCDE operon is problematic. Apparently, when growing on propionate itself, the uninduced levels of propionyl-CoA synthetase (PrpE) are not enough to generate sufficient propionyl-CoA to react with oxaloacetate and generate the inducer, methylcitrate (130). Consequently, acetyl-CoA synthetase (Acs), which can utilize either acetate or propionate as substrates, is also required for growth on propionate. Acs is needed to generate sufficient initial levels of propionyl-CoA to allow induction (130). Like acetyl-CoA synthetase (see above), the activity of propionyl-CoA synthetase (PrpE) is controlled by acetylation of the protein by the CobB sirtuin (130). Since functional Acs is needed for induction, the CobB protein is also required for expression of the prpBCDE operon during growth on exogenous propionate (171). However, in S. enterica induction of the propanediol utilization (pdu) operon by 1,2-propanediol abrogates the requirement for CobB. This is due to the presence in the pdu system of another sirtuin, which can functionally replace CobB (171).
It was noted long ago that E. coli requires an exceedingly long lag phase before growth on propionate proceeds (87, 169). Sequence analysis has shown that the E. coli prpBCDE operon contains four tandem 91-bp repeats inserted between prpB and prpC (86). It seems plausible that the 4- to 7-day adaptation time is due to selection of mutants lacking the insert rather than to slow genetic induction (87). Propionate is rather toxic and increased sensitivity is seen in mutant strains lacking the methylcitrate cycle. Toxicity appears to be largely due to the generation of 2-methylcitrate (74). In the absence of the PrpC 2-methylcitrate synthase, 2-methylcitrate can still be made by the citrate synthase of the citric acid cycle (GltA). Under these conditions gltA mutants having attenuated citrate synthase activities arise as propionate-resistant strains (74).
Aerobic growth on fatty acids results in their complete conversion to acetyl-CoA; consequently, the glyoxylate cycle plays the same essential role as when acetate is sole carbon source. The generation of one acetyl-CoA from each two-carbon segment of a fatty acid molecule results in the concomitant production of one equivalent each of FADH2 and NADH, which may be used in ATP generation. This ATP is not forthcoming during acetate (or acetoacetate) degradation and thus more ATP (on a per carbon atom basis) must be expended to activate these molecules. Hence growth on fatty acids is energetically more favorable (per carbon atom) than growth with either acetate or acetoacetate. Aerobic growth of wild-type E. coli K-12 on fatty acids as sole carbon sources occurs only when the fatty acid is 12 or more carbons long and then proceeds only after a distinct lag period required for induction of the fad regulon. Fatty acids of 6 to 10 carbon atoms may be utilized by E. coli cultures that have been preinduced with longer fatty acids or by strains in which expression of the fad regulon is constitutive due to mutations in the fadR gene (119, 120, 127, 128, 153, 179). For this reason we refer to fatty acids of >C12 as long-chain fatty acids and those with C6 to C10 as medium-chain fatty acids. Carboxylic acids shorter than C6 are referred to as short-chain fatty acids and cannot be metabolized solely via the fad system.
Most of our knowledge of the genetics and biochemistry of fatty acid degradation has been derived from studies with E. coli grown aerobically. The synthesis of at least five proteins involved in fatty acid β-oxidation (Fig. 6, Table 4) are coordinately induced when long-chain fatty acids are present in the growth medium. The genetic and enzymological studies of Peter Overath and coworkers constitute the basis of the more recent studies. The Overath group published the last of their papers on fatty acid catabolism over 30 years ago; their contributions were thoroughly reviewed by Nunn in the first edition of this book (120) and elsewhere (119). For these reasons we shall limit our discussion (except where necessary) to the developments in the field since the Nunn reviews and refer persons requiring the older literature to those reviews. However, note that the data and interpretations of the Overath publications have withstood the test of time in exemplary fashion. The genes encoding the enzymes of aerobic β-oxidation (the fad regulon) are scattered around the E. coli chromosome and fadBA are the only cotranscribed genes (note, however that there is some overlap with the newly discovered anaerobic degradation system; see below). The fad regulon is primarily responsible for the transport, acylation, and β-oxidation of medium-chain fatty acids (C7 to C11) and long-chain fatty acids (C12 to C18) and the expression of these genes is specifically controlled by the fadR gene product. In addition to enzymes of the fad system, growth of E. coli on short-chain fatty acids (C4 to C6) requires two degradative enzymes (Fig. 4) encoded by the atoD, atoA, and atoB genes and regulated by the atoC gene product.
Table 4Gene-protein relationships in fatty acid oxidation |
First, we will examine the genetic and enzymological information regarding the aerobic degradative pathways for both long- and short-chain fatty acids. We will then discuss the newly discovered anaerobic fatty acid utilization pathway and the mechanisms of fatty acid uptake. Finally, the mechanisms whereby the fad and ato structural genes are regulated will be considered.
The pathways by which E. coli degrades fatty acids (Fig. 6) are substantially similar to the β-oxidative pathways present in the mitochondria of mammals and other eukaryotic organisms. These pathways are classical examples of the oxidation of a series of homologous substrates through a series of homologous intermediates. With each turn of the β-oxidation cycle, the fatty acyl-CoA loses a two-carbon fragment as acetyl-CoA and reduces one molecule of flavin adenine dinucleotide (during the acyl-CoA dehydrogenase reaction) and one molecule of NAD (during the 3-hydroxyacyl-CoA dehydrogenase reaction). Acetyl-CoA, produced in the CoA-dependent thiolytic cleavage, is further metabolized in the tricarboxylic acid cycle. The other product of the cleavage step, the shortened fatty acyl-CoA molecule, reenters the degradation cycle without further activation.
The first step of fatty acid degradation is the activation of the free fatty acid to an acyl-CoA thioester by an acyl-CoA synthetase (fatty acid:CoA ligase [AMP forming]; EC 6.2.1.3). This reaction (Fig. 6) requires two high-energy phosphate equivalents per molecule of free fatty acid activated. The prediction of Overath and coworkers (90, 127) that aerobically grown E. coli cells contain a single acyl-CoA synthetase having broad specificity for medium- and long-chain fatty acids based on analysis of fadD mutants has been confirmed by purification of the enzyme to homogeneity. Although acyl-CoA synthetase was previously reported as a membrane-associated protein, Kameda and Nunn (85) found that over 90% of this enzyme was present in cytoplasmic fractions. Mangroo and Gerber (106) reported that the degree of membrane association of the acyl-CoA synthetase depends on the state of energization of the cell membrane. However, although a portion of FadD is clearly membrane bound (111, 127, 145, 186) various considerations argue that membrane association may not be physiologically relevant (see below). The molecular weight of the native enzyme was approximately 130,000 and the subunit molecular weight determined by polyacrylamide gel electrophoresis in the presence of sodium dodecyl sulfate was 47,000 (186). These experiments suggested that the enzyme may be a dimer or a trimer composed of identical subunits (24). The fadD gene has been cloned and sequenced (14). A single open reading frame encoding a protein with a molecular weight of 62,000 was found. The difference between this value and that observed by gel electrophoresis has recently been shown due to clipping of the enzyme in cell extracts by the OmpT outer membrane protease (186). The upstream regulatory region contained two operator sites for the fadR repressor as suggested by comparision to other fad regulon genes and directly confirmed by DNase footprinting (14). Strains carrying point mutations within the conserved acyl-CoA synthetase motif sequences show the expected decreases in activity (181).
In E. coli and S. enterica rather little is known about the enzyme responsible for the step following fatty acid activation, acyl-CoA dehydrogenase (Fig. 6). This reaction, the first in the cycle of degradation of the acyl chain, involves the transfer of two electrons from the substrate to an FAD cofactor that must be reoxidized for the dehydrogenase to have catalytic function. Indeed, the identity of the gene encoding this activity was only recently determined (26). Klein (89) reported the isolation of several E. coli mutants lacking functional acyl-CoA dehydrogenase and reported that these mutants mapped to at least two separate loci closely linked to proAB. The reported three-factor crosses and deletion data placed the fadE allele between proA and proB at min 5.62 (89). However, the subsequent E. coli K-12 genome sequence showed that only 12 bases lie between the two pro genes and thus these mapping data are clearly in error. Further confusion stemmed from the report of Klein (89) of two other acyl-CoA dehydrogenase mutants called fadF and fadG (although he suspected that these mutants might be allelic). These mutants were mapped between proA and the lac operon. However, these data were also in error since Δ(pro-lac) strains that carry deletions of this chromosomal segment grow well on fatty acids as sole carbon source, indicating the lack of essential fad genes within this region. Klein speculated that the fadF and fadG genes might encode proteins having different chain length specificities, similar to those found in the β-oxidation pathway of mammalian mitochondria and concluded that at least two genes are necessary for acyl-CoA dehydrogenase activity (89). He concluded that fadE comprised the electron transfer component of the acyl-CoA dehydrogenase reaction and that the products of genes fadF and fadG carried out the actual dehydrogenation reaction. The fadF and fadG strains are no longer available and no similar mutants have been isolated. The identification of yafH, an open reading frame of unknown function, as fadE, has recently resolved this situation. Mutants of both E. coli and S. enterica having disruptions of the yafH gene fail to grow on fatty acids aerobically (26, 157). Plasmids expressing yafH complemented growth on fatty acids of strains carrying fadE62, the best characterized of the Klein mutant alleles. In addition, the mutant allele was shown to contain a frameshift mutation (26, 157). It should be noted that a group working with S. enterica renamed yafH as fadF (158). However, this is incorrect because the complementation data demonstrate that yafH is fadE. No firm kinetic data are available for FadE since all the experiments reported by Klein involved adding purified enzyme to extracts of various fadE mutant strains (89); thus, further work is indicated.
The enoyl-CoAs produced by FadE serve as substrates for the cytosolic multienzyme FadBA complex, which has a broad chain-length specificity. The complex is a tetramer composed of two copies each of the 78-kDa α-subunit encoded by the fadB gene, and the 42-kDa β-subunit, the product of fadA (10, 11, 122, 134, 140). The two genes form an operon located at min 87 of the chromosomal map. This operon is negatively regulated by FadR and also subject to catabolite repression. Several groups have purified this complex, which has an overall molecular mass of 260,000 Da and found it to have an α2β2 structure (Fig. 7). Five enzyme activities, 3-ketoacyl-CoA thiolase, enoyl-CoA hydratase, L-3-hydroxyacyl-CoA dehydrogenase, cis-L-3-trans-L-2-enoyl-CoA isomerase, and 3-hydroxyacyl-CoA epimerase, are present in this multienzyme complex (10, 11, 122, 134, 140). The 3-ketoacyl-CoA thiolase activity is associated with the smaller subunit whereas the remaining four enzyme activities are associated with the larger 78-kDa subunit (10, 11, 122, 134, 140) (Fig. 6). The enoyl-CoA hydratase and 3-hydroxyacyl-CoA epimerase activities were found to utilize the same active site on the 78-kDa α-subunit. The cis-L-3-trans-L-2-enoyl-CoA isomerase activity is adjacent to the hydratase/epimerase but has distinct catalytic residues (185). However, it remains possible that the activities share a common CoA binding site. Such overlapping functional regions provide an explanation for the numerous activities resident in a protein of modest size. In saturated fatty acid oxidation, three final steps of the pathway are needed: enoyl-CoA hydratase, 3-hydroxyacyl-CoA dehydrogenase, and 3-ketoacyl-CoA thiolase. All are catalyzed by the complex (Fig. 6). When unsaturated fatty acids are degraded, two additional activities also carried by the complex, cis-2-enoyl-CoA isomerase and 3-hydroxyacyl-CoA epimerase, are required (Fig. 6).
Overath et al. predicted that 3-ketoacyl-CoA thiolase, 3-hydroxyacyl-CoA dehydrogenase, enoyl-CoA hydratase, and possibly 3-hydroxyacyl-CoA epimerase and cis-2-enoyl-CoA isomerase form an operon (127, 128). Their evidence was based on the highly coordinate induction of 3-ketoacyl-CoA thiolase, 3-hydroxyacyl-CoA dehydrogenase, and enoyl-CoA hydratase, as well as on the genetic-mapping results with mutants deficient in (i) all five enzymes (fadBA), (ii) 3-keto-acyl-CoA thiolase (fadA), and (iii) 3-hydroxyacyl-CoA dehydrogenase. This hypothesis was confirmed by cloning and analysis of the genes. The fadA gene was found to encode the β-subunit, whereas the fadB gene was found to be associated with the α-subunit. Sequencing and insertional mutagenesis of the fadBA operon showed that transcription proceeds from fadB to fadA rather than the opposite direction as originally reported (159). The reading frames of fadB and fadA are separated by only 10 nucleotides, suggesting possible translational coupling. Substantial similarities were found between the FadB and FadA proteins of E. coli and several of their eukaryotic counterparts involved in fatty acid β-oxidation. It seems that FadA may be either unstable or inactive in the absence of FadB. Plasmids carrying the fadA gene gave 3-ketoacyl-CoA thiolase I activity in fadA mutants but not in fadBA mutants. Moreover, fadA mutants carrying multicopy fadBA plasmids express both the 78- and 42-kDa proteins and give amplified levels of 3-ketoacyl-CoA thiolase activity. However, fadA mutants containing multicopy fadA plasmids have only wild-type levels of 3-ketoacyl-CoA thiolase activity despite expressing the 42-kDa fadA product from the plasmid (159). Yang and Schulz (183, 184) have found that the short-chain enoyl-CoA hydratase is associated with the fadBA multienzyme complex. They have argued that the product of this active site is channeled to the 3-hydroydecanoyl-CoA dehydrogenase active site also present on the FadB subunit without equilibration with the bulk solvent. Two additional enzymatic activities, cis-2-enoyl-CoA isomerase and 3-hydroxyacyl-CoA epimerase, possessed by the α-subunit are required when unsaturated fatty acids are degraded (155, 185). Recently it has been reported that a dead-end product formed by isomerization of the 2-trans-5-cis-tetradecenoyl-CoA intermediate of oleic acid oxidation to the 3,5-isomer is released from the enzyme complex by a thioesterase that may be the enzyme encoded by the tesB gene (142).
Another gene in the fad regulon, fadH, is located at min 71 to 75 on the E. coli chromosome far away from all previously known fad loci. The fadH gene encodes a 2,4-dienoyl reductase that is required only for the degradation of unsaturated fatty acids whose double bond extends from an even-numbered carbon atom (187). Mutants with lesions in fadH were isolated as unable to use petroselinic acid (cis-6-octadecenoic acid) but capable of growth on oleic acid (cis-9-octadecenoic acid). FadH is a rather unusual enzyme in that it contains three cofactors, both FMN and FAD plus a [4Fe-4S] cluster, as shown by a recent crystal structure (77).
Although the aerobic utilization of fatty acids in E. coli is now reasonably well understood, the environment in which this organism seems likely to encounter appreciable supplies of fatty acids, the mammalian gut, is thought to be severely limited in oxygen. Hence, although the intestinal lumen may be rich in a variety of fatty acids, utilization of these carbon sources by the aerobic pathway seems unlikely to occur. However, alternate electron acceptors such as nitrate, fumarate, and trimethylamine oxide that support anaerobic respiration may be available in the lumen. For these reasons Campbell and coworkers tested the ability of E. coli K-12 to grow anaerobically on fatty acids in the presence of nitrate, fumarate, or trimethylamine oxide and readily obtained growth (28). This was contrary to the implications of the report of Iuchi and Lin (79) that expression of 3-hydroxyacyl-CoA dehydrogenase and, to a lesser extent, acyl-CoA dehydrogenase activity, as well as major enzymes of the citric acid cycle and glyoxylate cycle, were repressed under anaerobic conditions via the ArcAB system.
The new anaerobic fatty acid utilization pathway is distinct from the aerobic pathway in that (i) it proceeds normally in mutant strains lacking various enzymes of the aerobic pathway (FadD, FadBA, and FadE), (ii) it functions with fatty acids (octanoate and decanoate) that cannot be utilized by wild-type E. coli strains under aerobic conditions, and (iii) superrepressor mutants of the fadR regulatory locus that block aerobic growth on fatty acids fail to block the anaerobic pathway (28). The pathway proceeds by cleavage of the carbon chain between the β- and γ-carbons (carbon atoms 2 and 3) as in the classical β-oxidation pathway (Fig. 6) and produces acetyl-CoA as shown by the finding that growth requires a functional glyoxylate cycle (28). The ability of several fad mutant strains blocked in aerobic growth on fatty acids to grow anaerobically on fatty acids as a sole carbon and energy source in the presence of an alternative electron acceptor indicated that E. coli possesses an alternative set of enzymes for utilizing fatty acids under these conditions. The first of these alternative enzymes found were those encoded by the yfcYX genes. These genes were identified as encoding putative β-oxidation enzymes by sequence analyses (68) and subsequently by microarray results showing induction of their expression by fatty acids (25). The existence of these genes seemed likely to explain the ability of strains having fadA::Tn10 or fadB null mutations to grow slowly on fatty acids despite the presence of inactivating mutations. Indeed, the residual growth of these strains on fatty acids was eliminated by deletion of the yfcYX genes (28). The proteins encoded by the yfcYX genes were shown to have the enzyme activities postulated by Snell and coworkers (156). YfcX possesses both the enoyl-CoA hydratase and 3-hydroxyacyl-CoA dehydrogenase activities, whereas yfcY encodes thiolase activity. Transcription of the yfcYX genes was regulated by FadR. For these reasons these genes have been renamed as fadJ (replaces yfcX) and fadI (replaces yfcY). The work of Snell and coworkers (156) was focused on using β-oxidation to provide acyl chains for the synthesis of polyhydroxyalkanoates, which are commercially important biopolyesters ("bioplastics"). E. coli does not normally make appreciable quantities of such compounds. However, upon expression of the appropriate genes from polyhydroxyalkanoate-producing bacteria in E. coli, high levels of these compounds are made. Although not covered here, a sizable literature exists on the use of E. coli to make such products.
Expression of both the fadBA and fadIJ operons is regulated by FadR and thus it might be expected that the presence of a FadR superrepressor mutation would prevent anaerobic growth of an otherwise wild-type strain on oleate plus nitrate. However, this was not the case (28). Loss of activity of either FabBA or FadIJ in this strain background resulted in an inability to grow under these conditions, indicating that under repressing conditions where activities are low, both multienzyme complexes are required for growth. This could be due to a simple additive effect of the two complexes, but given the levels of FadR repression previously reported (10- to 15-fold) for the two enzymes (25), slow growth rather than the robust growth observed was expected. For this reason a model in which the substrate specificities of the two complexes complement one another to achieve more efficient utilization of fatty acid chains was favored. It is energetically more favorable to produce a given amount of acetyl-CoA by complete degradation of an acyl chain rather than through partial degradation of multiple acyl chains. This is because each acyl chain requires activation by ATP hydrolysis to enter the initial cycle of β-oxidation, whereas subsequent cycles require no further activation. Klein et al. (90) reported that aerobic cultures of wild-type cells (where FadBA is the dominant enzyme complex) perform incomplete oxidative degradation of fatty acid chains. These workers assayed the release of 14C from oleate labeled at various positions within the acyl chain. Relative to the carboxyl carbon, release of carbon atoms 10 and 18 were only 0.4 and 0.33, respectively. Therefore, an accumulation of short- and medium-chain-length intermediates would be expected in these cells. Klein and coworkers also examined the pattern of carbon release in a strain (fad-5) that lacks all of the fadBA-encoded enzyme activities. From the complete lack of aerobic growth of fadBA fadIJ double-mutant strains on fatty acids it seems very probable that the remaining β-oxidation activity reported in the fad-5 strain is that of the FadIJ complex. Cultures of the fad-5 strain released the distal carbons at much slower rates than the wild-type strain. Relative to the carboxyl carbon, the rates of release of carbon atoms 10 and 18 were only 0.07 and 0.03, respectively. Therefore it seems that the FadBA complex degrades long-chain fatty acids with fair efficiency, but releases appreciable quantities of short- and medium-chain-length intermediates that are excellent substrates for the FadIJ complex. In support of this hypothesis, fadIJ null mutant strains failed to grow anaerobically on the shorter-chain fatty acids, indicating that anaerobic growth on short- or medium-chain fatty acids requires FadIJ activity. This "efficient utilization" model provides a rationale for retention of the two β-oxidation complexes. The results of Snell and coworkers (156) somewhat disagree with this model. In vitro the enzyme activities assayed by these workers had a preference for medium-chain rather than short-chain substrates. However, only a limited range of substrates was examined, which did not include long-chain substrates, and thus further analyses are needed to test the efficient utilization model. Although the fadIJ operon is regulated by FadR, both aerobically and anaerobically, anaerobic transcription of these genes appears to involve factors in addition to FadR. The fact that wild-type strains are capable of growth on medium-chain fatty acids anaerobically indicates that FadR does not play a major role in regulation of anaerobic fatty acid degradation.
In addition to enzymatic activities targeting the β-carbon that result in thiolytic cleavage of the chain a β-oxidation pathway requires enzymatic activation of free fatty acids to their CoA thioesters (which facilitates chemistry at the β-carbon) and redox protein(s) that link the reducing equivalents produced by β-oxidation to the respiratory chain. These activities appear likely to be provided by the ydiQRSTD operon, although the function of only the last gene of the operon, ydiD (now named fadK), has been demonstrated (111). FadK is an acyl-CoA synthetase having preference for short-chain-length fatty acid substrates (<10 C atoms). The enzymatic mechanism of FadK is similar to other acyl-CoA synthetases in that it forms an acyl-AMP intermediate prior to the formation of the final acyl-CoA product (111). Expression of FadK is repressed during aerobic growth and is maximally expressed under anaerobic conditions in the presence of the terminal electron acceptor, fumarate. Either the FadD or FadK proteins can support anaerobic oleic acid utilization in the presence of nitrate (111).
The putative ydiQRSTD operon encodes several proteins (in addition to FadK) that could play a role in anaerobic β-oxidation. Proceeding upstream from fadK, the genes encode a putative ferrodoxin (YdiT), a flavoprotein (YdiS) that could accept electrons and use these to reduce a quinone and two proteins (YdiR and YdiQ) that seem likely to form a heterodimeric electron transport flavoprotein (that could transfer electrons to YdiS). There is also a putative acyl-CoA dehydrogenase, ydiO, located immediately upstream of the ydiP gene (which is immediately adjacent to ydiQ). The ydiQRST genes have high sequence homology to the fixABCX operon of anaerobic carnitine metabolism and may function as an electron transport chain linking the β-oxidation of fatty acids with the anaerobic respiratory chain. Most of these genes would be involved in electron transfer from the multienzyme complexes under anaerobic conditions since FadE performs this function under aerobic conditions (26, 90). However, since fadE null mutant strains grow normally on oleate plus nitrate under anaerobic conditions (28), E. coli must possess another route of electron transfer from the multienzyme complexes, and the products of the ydiOQRST genes seem prime candidates for this function. This model resembles fatty acid metabolism in mammalian mitochondria where several electron transfer flavoproteins link β-oxidation of fatty acids with the respiratory chain. Together with fadIJ the genes of this putative operon could form a complete alternative β-oxidation system.
Prior to catabolism, fatty acids must enter the cell via uptake systems that translocate these hydrophobic compounds across both the outer and inner membranes. Prior to kinetic analyses the prevailing thought was that fatty acids diffuse through membranes without requiring a protein carrier. This is probably true for medium- and short-chain fatty acids where no transport proteins or genes required for transport have been identified. However, in the case of long-chain fatty acids physiological and kinetic studies from several laboratories (16, 33, 44) indicate that a carrier mechanism facilitates their entry into E. coli. Furthermore, in addition to the FadD and FadK acyl-CoA synthetases the product of the fadL gene is required to deliver exogenous long-chain fatty acids across the cell membrane to the cytosolic fatty acid degradative enzymes (Fig. 8). The fadL gene encodes for a 43,000-Da outer membrane protein essential for long-chain fatty acid transport and is the paradigm protein of a conserved family of outer membrane proteins involved in the uptake of hydrophobic compounds (175).
The FadL protein was originally reported to be located in the inner membrane (61). However, Morona and Henning (112) found that mutants in the ttr locus abolished growth on long-chain fatty acids as well as conferring resistance to bacteriophage T2 and that ttr was genetically indistinguishable from fadL. Subsequent work by Black (12) showed that ttr and fadL are identical and that the FadL protein is indeed an outer membrane protein as indicated by its function as a phage receptor (12). The FadL protein was heat modifiable, a characteristic of outer membrane proteins. The fadL gene encodes a protein of 448 amino acid residues with a molecular weight of 48,831. Maturation removes a signal peptide of 27 amino acids, leaving a native protein of 421 residues and a molecular weight of 45,969 (13). A native molecular mass of approximately 130,000 Da (13) suggested a trimeric structure reminiscent of that found for OmpC, OmpF, and other porins. Therefore, FadL seemed likely to be a porin specific for long-chain fatty acids analogous to the LamB maltodextran-specific porin of maltose catabolism. However, a recent crystal structure (175) shows FadL to be quite different from LamB and other trimeric proteins in that it is monomeric rather than trimeric and is a 14-stranded beta barrel, the opening of which is occluded by a central hatch domain. The structures suggest that hydrophobic compounds bind to multiple sites in FadL and use a transport mechanism that involves spontaneous conformational changes in the hatch. In this view, FadL would use a transport mechanism that is based on diffusion coupled with spontaneous conformational changes and the major function of the protein would be to allow fatty acids to efficiently cross the outer membrane without being partitioned into the hydrophobic lipid layers of the membrane. Binding of fatty acids to FadL occurs at the C-terminal end of the protein and single-residue changes eliminate fatty acid binding (94).
The FadL protein was specifically labeled using a photoaffinity label attached to a fatty acid, whereas no fatty acid binding protein was detected in the inner membrane by using this approach (106). Although it remains possible that the putative inner membrane carrier failed to bind the photoaffinity-labeled fatty acid (which was derived from a medium-chain fatty acid), the observed lack of labeling is consistent with the idea that fatty acids cross the cytoplasmic membrane by a simple diffusion mechanism (106). Some further evidence in favor of this idea is that both enantiomers of lipoic acid (which can be considered a modified octanoic acid) are taken up by E. coli, although only the R-lipoic acid becomes attached to the 2-oxoacid dehydrogenases (123). Since a protein transporter would be expected to discriminate between enantiomers, this finding argues against the existence of a transporter for lipoate or for fatty acids of similar chain length. Higashitani et al. (71) made an interesting observation that expression of FadL is regulated in response to osmolarity. Transcription of the fadL gene is repressed at high external osmotic pressure and transport of long-chain fatty acids is abolished under these conditions. Osmoregulation of fadL is due to binding of OmpR protein to four sites around the fadL promoter, and thus fadL seems subject to the OmpR/EnvZ two-component regulatory system (71).
Several observations demonstrate that the long-chain fatty acids enter E. coli via an active unidirectional mechanism. (i) The transport of long-chain fatty acids into wild-type strains is a saturable process implying the existence of a binding protein (104, 121). (ii) No efflux of transported radioactive long-chain fatty acids occurred when cells of wild-type E. coli strains were washed with unlabeled long-chain fatty acids (104, 121). (iii) Both the energy of activation and the Q10 of long-chain fatty acid transport are representative of enzyme-mediated processes (104). As indicated above, both an acyl-CoA synthetase and FadL are required to deliver long-chain fatty acids across the membrane to the cytosolic fatty acid-degradative enzymes of E. coli. Since FadL does not require energy and the protein is located in the outer membrane, the active component required to deliver long-chain fatty acids to the cytosol appears to be the acyl-CoA synthetase.
Many years ago Overath predicted that fatty acid transport would proceed by vectorial thioesterification (127). The attachment of CoA would trap the fatty acids in the cytosol and thereby account for overall unidirectional uptake. Indeed Overath found that fadD mutants are unable to accumulate exogenous fatty acids of any length into either the cytosol or the membrane lipids of E. coli (127). Recent experiments have shown that long-chain fatty acid translocation in E. coli membrane vesicles requires both acyl-CoA synthetase and ATP and that acyl-CoAs accumulate (151).
In principal the physical transfer of fatty acids across the cytoplasmic membrane, even for long-chain fatty acids, does not require any energy or the involvement of any protein. It has been argued (69, 91) that nonionized long-chain fatty acids can transit a phospholipid bilayer by a flip-flop process with a half-life <2 s, a rate sufficiently fast for effective transport in small cells. In contrast, ionized fatty acids cross the bilayer much more slowly (half-life of several minutes). Although membrane association of FadD has been proposed to play a role in fatty acid transport (106), the fact that FadD can be functionally replaced by an acyl-CoA synthetase from yeast (92) makes this seem unlikely. An oleic acid binding protein distinct from FadD was partially purified from cell envelopes and characterized (84). This protein has been postulated to be an essential inner membrane H+/fatty acid transporter (86), but no genetic or further biochemical data supports the existence of this component. Therefore the present data argue strongly that transmembrane movement of fatty acid across the inner membrane of E. coli occurs by a passive diffusion vectorial thioesterification mechanism. A second fatty acid transport system of much lower capacity (ca. 2% of the FadD pathway) is also found in E. coli. This FadD-independent (but FadL-dependent) transport was detected by the incorporation of labeled fatty acids into membrane lipids and is due to an acyl-ACP synthetase that incorporates of fatty acids into membrane phospholipids via their acyl-ACP derivatives (80, 144). However, this is a tightly coupled system that seems unable to deliver long-chain fatty acids to the cytosol and can be considered a second very specialized example of vectorial thioesterification.
Mutants lacking the FadE, FadA, or FadB enzymes have decreased transport rates for medium- and long-chain fatty acids relative to wild-type strains (58, 90). These findings imply that fatty acid transport is coupled to fatty acid oxidation. Although the mechanism(s) is unknown, it is conceivable that transport is reduced in these fad mutants as a consequence of feedback inhibition. For example, the accumulation of fatty acyl-CoA intermediates in such fad mutants may be inhibitory to acyl-CoA synthetase. Nonetheless, β-oxidation is not an absolute requirement for transport because unsaturated fatty acid auxotrophs carrying a fadE mutation incorporate exogenous unsaturated fatty acids into phospholipid (90).
The degradation of acetoacetate to acetyl-CoA was also first elucidated by Overath and coworkers (133), who showed that the pathway was composed of two steps (Fig. 4). First, acetoacetate is activated to acetoacetyl-CoA in a reaction catalyzed by acetyl-CoA:acetoacetyl-CoA transferase. The acetoacetyl-CoA is then cleaved to two molecules of acetyl-CoA by 3-ketoacyl-CoA thiolase II (55, 56, 133, 160, 161). These enzymes are highly inducible by acetoacetate and are specific for β-keto derivatives of short-chain acyl-CoAs. The earliest studies identified the loci responsible for acetoacetate degradation as two closely linked genes: atoA encoding acetoacetyl-CoA transferase and atoB encoding 3-ketoacyl-CoA thiolase II. Subsequent biochemical work showed that acetoacetyl-CoA transferase is a heterotetrameric protein composed of two α− and two β-subunits (57, 160), whereas 3-ketoacyl-CoA thiolase II is a homotetrameric protein (49). These results indicated that a second acetoacetyl-CoA transferase gene encoding the second subunit must exist. Molecular cloning of the known gene cluster showed that this missing gene (called atoD) was linked to the other ato genes and that the three structural genes comprise an operon transcribed from atoD to atoB (Fig. 3). Acetoacetate serves as metabolic inducer for the ato system. When acetoacetate is used as sole carbon source, a 200- to 300-fold induction of both acetoacetyl-CoA transferase and 3-ketoacyl-CoA thiolase II is seen. Mutants deficient in 3-keto-acyl-CoA thiolase (fadA), or 3-hydroxyacyl-CoA dehydrogenase plus enoyl-CoA hydratase (fadB) activities have decreased transport rates for medium- and long-chain fatty acids relative to wild-type strains (34, 44). These findings imply that fatty acid transport is coupled to fatty acid oxidation. Although constitutive levels of the three required fad system enzymes are present in fadR strains, no growth on the saturated C4 or C5 acids occurs because these substrates fail to induce the ato enzymes (81, 82, 133). Only strains having constitutive levels of the needed ato- and fad-encoded enzymes utilize C4 or C5 acids as sole carbon source. Overath and coworkers first showed that mutants (called But+ mutants) that grow on butyrate or valerate are readily isolated by plating fadR mutants on minimal medium containing butyrate (133). Most such But+ mutants are constitutive for the ato enzymes because of mutations in the atoC regulatory gene (81, 82). Hence, strains that utilize C4 and C5 as sole carbon sources have the genotype fadR atoC.
Acetoacetyl-CoA transferase encoded by the atoDA genes is the only component required for uptake of short-chain fatty acids. There is no evidence for any outer membrane transport component. The short-chain fatty acids presumably cross the outer membrane via porin channels and diffuse across the cytoplasmic membrane in the nonionized form. Once they enter the cytosol the short-chain fatty acids are converted to their CoA thioesters by acetoacetyl-CoA transferase in a trapping mechanism similar to that played by FadD and FadK in long- and medium-chain fatty acid uptake. Acetoacetyl-CoA transferase utilizes acetoacetate as well as C4-, C5-, and, perhaps to some extent, C6-saturated carboxylic acids (160, 161, 162). Frerman and Bennett (56) showed that membrane vesicles prepared from cells induced by acetoacetate translocated C4 acids. Uptake was stimulated by ATP and acetyl-CoA and did not occur in membrane vesicles from noninduced cells. Frerman and Bennett (56) also found that significant amounts of acetoacetyl-CoA transferase were associated with the membrane and that uptake was rapidly inhibited by butyryl-CoA and acetate, the products of the reaction catalyzed by acetoacetyl-CoA transferase (Fig. 4). Although these results show the importance of acetoacetyl-CoA transferase in short-chain fatty acid transport, the possibility that other components play a role is not excluded. Mutant studies have showed that atoD mutants and also atoB and fadBA (in the case of butyrate) mutants have reduced short-chain fatty acid transport, suggesting a coupling between metabolism and transport similar to that observed for long-chain fatty acids (133).
The fatty acid degradation (fad) system is primarily responsible for the transport, activation, and aerobic β-oxidation of medium-chain (C7 to C11) and long-chain (C12 to C18) fatty acids. Wild-type E. coli strains grow using long-chain (>C12) fatty acids as sole carbon and energy source. However, these strains are unable to grow using medium- or short-chain fatty acids. Although the enzymes of β-oxidation function on the shorter-chain-length substrates, such fatty acids are unable to induce the genes of the fad system. Consequently, growth on medium- or short-chain fatty acids requires constitutive expression of the E. coli fad regulon. In contrast, many bacteria (e.g., pseudomonads) grow on fatty acids of all chain lengths and it has been shown that, in these cases, fatty acids of C6 or longer are effective inducers of the β-oxidation system (148). The fad structural genes, which map at six distinct loci on the E. coli chromosome are regulated by the fadR gene (39, 127). When consecutive cycles of β-oxidation have shortened a fatty acid to the four-carbon stage, the acetoacetyl-CoA induces the ato operon whose products are required for the final step of converting this intermediate to two molecules of acetyl-CoA. Growth on saturated short-chain fatty acids requires the action of enzymes of the fad system and also operation of the ato system (see above).
The levels of the enzymes of aerobic fatty acid degradation in E. coli depend on at least three regulatory systems, the global Crp/cyclic AMP and ArcAB systems plus the fatty-acid-specific fadR gene. The Crp/cyclic AMP system exerts its classical positive control of carbon utilization (34, 132) (putative Crp binding sites are found in the promoter regions of the fad genes), whereas FadR negatively controls both the fad regulon (26, 28, 45, 70, 127, 128) and the aceBA operon (65, 105). The ArcAB system has been shown to strongly (>20-fold) repress expression of the 3-hydroxyacyl-CoA dehydrogenase encoded by the fadB gene and weakly repress acyl-CoA dehydrogenase activity (79). The mechanisms of repression of these genes by the ArcAB system have not yet been explored but are likely to be the same as for the pathways into which the acetyl-CoA produced by β-oxidation flows, i.e., the glyoxylate and citric acid cycles. The FadR protein has a dual role in fatty acid metabolism as a repressor of the β-oxidation pathway and an activator of the fatty acid biosynthesis pathway (27, 39). In the fad regulon FadR acts as does LacI in the lac system; it is a classical transcriptional repressor (see below). The fadR gene was discovered by selection for mutants able to utilize medium-chain fatty acids by plating wild-type cells onto minimal medium containing decanoate as sole carbon source. This selection readily gives loss-of-function fadR mutants. The rationale for the selection was based on early studies showing that wild-type strains could switch to decanoate after first being grown on oleate, but continued for only a few doublings before growth ceased. Hence it was clear that decanoate was a carbon source, but not an inducer. Overath's original hypothesis (127) that the fadR gene product was a diffusible repressor protein has been validated by several lines of evidence. First, transposon-generated fadR null mutations confer constitutive expression of β-oxidation (153). Second, genetic studies with strains merodiploid for the fadR gene showed that the wild-type fadR allele is trans-dominant to fadR (154). Moreover, fatty acid oxidation in fadR strains harboring multicopy plasmids carrying wild-type fadR was decreased relative to wild-type strains containing one chromosomal copy of the wild-type fadR gene (although such strains remain inducible) (48). A third line of evidence is the isolation of genetically dominant fadR mutants that superrepress β-oxidation (78). Mutants that are refractory to inducer are expected of LacI-like repressors.
The first evidence that the fadR gene product regulates the expression of the fad regulon at the level of transcription was the lacZ transcriptional fusion studies of Clark (34). LacZ expression in strains carrying such fusions was inducible by long-chain fatty acids in wild type strains and constitutive in fadR strains. Furthermore, the expression of β-galactosidase was repressed in these strains under catabolite-repressing growth conditions (as are the levels of the fatty acid degradative enzymes) and overexpression of FadR gave increased repression, presumably because of increased occupation of operator sites. Overath found that the β-oxidation enzymes could not be induced in fadD mutants, whereas mutants with lesions in the other fad genes remained inducible (90). He postulated that long-chain acyl-CoA thioesters are the in vivo inducers of the fad regulon. In vitro studies confirmed this postulate. DiRusso et al. (46) demonstrated binding of purified FadR protein to a fragment from the fadBA promoter and localized binding to a region just downstream of the transcriptional start site. Furthermore, FadR binding inhibited fadBA transcription in an in vitro system composed of purified components. DNA binding by FadR is inhibited by long-chain fatty acyl-CoAs consistent with the Overath proposal. The Ki values found were approximately 5 nM for acyl-CoAs with C16 or C18 fatty acids, whereas myristoyl-CoA showed a Ki of 250 nM. Decanoyl-CoA and free fatty acids inhibited binding only at millimolar concentrations. Later work also demonstrated binding to the promoter regions of the fadL and fabA genes (47). The affinity of binding was greatest for fadBA, next for fadL, and least for fabA (where FadR acts as an activator, not a repressor; see below). A shortcoming of the first report (46) that long-chain acyl-CoAs release FadR from specific DNA binding sites was the failure to control for the fact that long-chain acyl-CoAs are strong ionic detergents and hence can be powerful protein denaturants akin to sodium dodecyl sulfate. Thus, the observed dissociation could have been due to protein denaturation rather than the postulated specific ligand-protein interaction. These workers used nonionic detergents in an attempt to control for detergent action, but nonionic detergents are very weak protein denaturants and thus this control was not convincing. However, Henry and Cronan (70) showed that acyl-CoA-mediated release of FadR from the fabA operator was fully and rapidly reversed by thioesterase cleavage of the acyl-CoA. Since detergent denaturation of proteins is characteristically reversed only extremely slowly and inefficiently, this result coupled with the low levels of long-chain acyl-CoA needed for release plus the correlation of the acyl-CoA chain-length dependence with the fatty acid induction specificity observed in vivo provided strong evidence that acyl-CoA is the fad regulon inducer. However, it remained possible that some other fatty acid derivative was the inducer in vivo. The observation of Klein et al. (90) that fadD mutants were noninducible fell short of providing evidence that acyl-CoAs were the in vivo inducers because fatty acids fail to enter these cells. However, Cronan (38) showed that fatty acids diverted from the fatty acid synthesis pathway induce the fad regulon, but only when the cells contained functional FadD, thereby showing that acyl-CoAs, rather than fatty acids, are the inducer.
Henry and Cronan (70) showed that the FadR protein activates transcription of a gene (fabA) that plays a key role in fatty acid biosynthesis in E. coli. The fabA gene is largely repressed in vivo by the presence of exogenous fatty acids. (The function of FadR as a positive activator will be discussed more fully in the chapter on lipid biosynthesis.) FadR exerts different effects on the synthesis and degradation of fatty acids based on the location of the binding site on the DNA. In the fabA case the binding site is upstream of the promoter elements at the location where activator proteins typically bind to assist RNA polymerase action. In contrast the binding sites for the genes of the fad regulon lie within the promoter where binding of regulatory proteins impedes RNA polymerase. The sequences to which FadR binds are weakly palindromic and have a consensus of 5'-TGGTNNNACCA-3' (182). The fadL gene has two such sequences, although expression of this gene appears to be only weakly (<twofold) regulated by FadR. Expression of fadD is likewise weakly regulated as expected from the fact that FadD and FadL are needed to produce the intracellular inducer. Moreover, the sequence upstream of the fadL gene also contains a putative Crp binding site, although glucose fails to repress expression of the gene. Indeed, a difficulty in our understanding of the regulation of β-oxidation is that the levels of induction observed for β-oxidation in E. coli vary markedly among different reports. The variation observed for a given medium suggests that a regulatory system other than FadR is complicating the results. A likely possibility is anaerobic regulation mediated by the ArcAB system which could vary with the cell density and oxygenation of putatively aerobic log-phase cultures, and thus it would seem advantageous to assay FadR action in strains defective in ArcAB regulation.
Crystal structures of uncomplexed E. coli FadR as well as for cocrystals of FadR with operator DNA or with tetradecanoyl-CoA have recently been reported (173, 174, 182). FadR is a two-domain molecule where the N-terminal domain binds DNA and the C-terminal domain binds acyl-CoA. The structure of the N-terminal domain conforms to the ‘winged-helix’ motif consistent with its role in binding DNA. The larger C-terminal domain is essentially an antiparallel array of seven helices that form a barrel-like structure, with an eighth helix forming a lid at the end (173, 174, 182). FadR forms a dimeric structure in the crystal, which is consistent with studies showing that the protein is functional as a dimer in solution (47). The current model of FadR action based on the crystal structures is that upon DNA binding the operator DNA is bent about 20° and five different regions of each FadR monomer contact the operator DNA. Each recognition helix makes several direct contacts with the bases of the major groove and one of the wings makes contacts deep in the minor groove where it binds two bases (the two G/C base pairs in each half-site). Long-chain fatty acyl-CoA binding results in a progressive series of large helix movements in which helix 8 becomes kinked upon binding acyl-CoA. The kinked helix-8 then pushes on helix-4 which in turn pushes on helix-1 of the DNA binding domain. The overall result is that the recognition helices are pushed apart by 7 Å and rotated by 13 Å, thereby preventing specific DNA binding. Essentially the energy of acyl-CoA binding provides a push that releases FadR from DNA.
The pathways for the uptake and aerobic degradation of fatty acids and acetoacetate are well investigated and their overall regulation is also largely understood except for minor details. Regarding degradation, the breakdown of fatty acids that generate "awkward" intermediates may yet provide a few more novel enzymes and their corresponding genes, as in the case of the fadH-encoded dienoyl reductase. As far as we know, no systematic study of the ability of E. coli to degrade unsaturated, acetylenic, hydroxylated, branched, or ring-containing fatty acids has yet been made.
One recent development has been the discovery of a pathway for fatty acid degradation that is expressed only under anaerobic conditions. There is much to be learned of the regulation and mechanism of this pathway. The analysis of such duplicate pathways that are used under different environmental conditions has been greatly aided by the availability of the E. coli chromosomal sequence. In particular, elucidation of the pathways for using "lesser" carbon sources, including glycolate, glyoxylate, and propionate, has greatly benefited from gene sequence comparisons.
Despite considerable knowledge of most aspects of their regulation, the induction of both acetyl-CoA synthetase and the glyoxylate operon in response to acetate are still puzzling. In particular the identity of the inducer molecule has remained elusive as has the role of the presumed regulatory gene, arpA. Moreover, the reversible acetylation of the acetyl-CoA synthetase protein and other targets by the sirtuin-related system (CobB and Pat) has opened up a new area of regulation.
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Module3.4.4
