Biotin and Lipoic Acid: Synthesis, Attachment, and Regulation
JOHN E. CRONAN
[SECTION EDITOR: T. BEGLEY]
January 2, 2008
Departments of Microbiology and Biochemistry, University of Illinois, Urbana, IL 61801
Mailing address: Department of Microbiology, B103 Chemical and Life Sciences Laboratory, University of Illinois, 601 S. Goodwin Ave., Urbana, IL 61801. Phone: (217) 333-7919, Fax: (217) 244-6697, E-mail:
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Biotin {vitamin H, vitamin B7, or 5-[(3aS,4S,6aR)-2-oxo-1,3,3a,4,6,6a-hexahydrothieno[3,4-d]imidazol-4-yl]pentanoic acid} and lipoic acid {5-[(3R)-dithiolan-3-yl]pentanoic acid, also called 6,8-thioctic acid} share many similarities (Fig. 1). Both vitamins are essential for aerobic growth of Escherichia coli and Salmonella enterica, whereas biotin is also required for growth of these bacteria under anaerobic conditions. Both biotin and lipoic acid must be covalently attached to their cognate proteins to perform their roles in cellular enzymology; the free vitamins are not physiologically useful (although free biotin plays an indirect regulatory role). The protein domains to which biotin and lipoic acid are attached have very similar three-dimensional structures and the enzymes that perform the attachment of the two cofactors are members of the same protein family based on their structures. Thus, the speculation made many years ago (212) that biotin and lipoic arose together "late" in evolution is germane. Moreover, although the two molecules appear to have little similarity when drawn as in Fig. 1, both are chiral. Biotin has a chair shape due to the C–N bonds, whereas the ring of lipoic acid is skewed by the C–S bonds. Proteins recognize these structures in somewhat similar manners since the biotin binding protein, avidin, also shows significant (albeit much weaker) binding of lipoic acid, and antibodies raised against one of the molecules as a hapten often bind to proteins modified with the other cofactor (84).
Biotin and lipoic acid also share the property that they are attached to very few protein species. E. coli has only a single biotinylated protein, the AccB subunit of the essential enzyme, acetyl-CoA carboxylase, whereas S. enterica has a second inducible biotinylated protein, the α-subunit of oxalacetate carboxylase (221, 225, 226). E. coli and S. enterica have three lipoylated proteins, these are subunits of pyruvate dehydrogenase (PDH) and 2-oxoglutarate dehydrogenase (2-OGDH), enzymes essential for aerobic growth, plus a third lipoylated protein induced by the presence of glycine that is a subunit of the glycine cleavage system of single carbon metabolism (211, 223, 224). In each of these proteins the cofactor is attached to a lysine residue ε-amino group of a domain of highly conserved structure. This domain is the N-terminal part of a lipoylated protein and the C-terminal part of a biotinylated protein and is connected to the remainder of the protein by a long proline plus alanine-rich linker region (159). The modified subunits form noncovalent interactions with other members of a protein complex of the three or four protein species that constitute the active enzyme. The cofactor-modified domains then shuttle intermediates between the multiple active sites of the enzyme complex (159). The mobility of the domains is due to the proline-alanine linkers and the domains constitute the distal ends (the "hands") of the swinging arms long ago postulated for these enzyme complexes. These arrangements can be considered to provide substrate channeling via covalent attachment (159). Finally both biotin and lipoic acid are needed in only trace quantities. In E. coli only a few hundred molecules of biotin per cell are sufficient for growth (51) and the requirement for lipoic acid is similar. Therefore, the enzymes of these pathways are expressed at very low levels (<350 molecules per cell; reference 133) and the enzymes have generally low turnover numbers.
The early steps of biotin biosynthesis are not well understood in any organism, but clearly differ between E. coli and Bacillus subtilis (although other bacilli seem to follow the E. coli pathway). In both cases a seven-carbon dicarboxylic acid, pimelic acid, is assembled with one of its carboxyl groups in thioester linkage. Pimeloyl-CoA has long been thought to be the thioester-activated form of pimelic acid, but recent evidence suggests a role for the acyl carrier protein (ACP) of fatty acid synthesis as the thiol moiety (192; S. Lin and J. E. Cronan, unpublished data). In contrast, the steps that follow formation of the pimeloyl-thioester are well conserved throughout biology even in organisms (e.g., Saccharomyces cerevisiae) that lack the ability to perform any early biosynthetic steps. In E. coli the atoms of biotin are derived from rather disparate sources: acetate, alanine, CO2, S-adenosylmethionine (SAM), and sulfide. Two groups have traced the origins of the biotin and dethiobiotin carbon atoms by 13C labeling followed by analysis by 13C NMR (105, 182). Using the numbering system of Fig. 1, the C-3, C-5, and C-7 carbon atoms of biotin are derived from the C-1 of acetate, whereas the C-2 of acetate contributes the biotin C-2, C-4, and C-6 carbon atoms. Acetate labeled in both carbon atoms is incorporated intact as shown by 13C coupling. Biotin carbon atoms C-9 and C-10 are contributed by l-alanine. The C-1 and ureido (C-2') carbon atoms are derived from CO2 (105). The nitrogen atom adjacent to C-7 is from SAM, whereas the other nitrogen atom is from alanine. The labeling pattern is consistent with the formation of a pimelic acid moiety by head-to-tail incorporation of three intact acetate units, as is the case in fatty acid (or polyketide) synthesis (105, 182), and the labeling pattern eliminates other plausible pathways from tryptophan, lysine, and diaminopimelic acid or elongation of 2-oxoglutarate (182). Moreover, the 13C labeling results eliminate free pimelic acid as an intermediate in biotin biosynthesis. Pimelic acid is a symmetrical dicarboxylic acid whose carboxyl groups cannot be stereochemically distinguished and if free pimelic acid was an intermediate, then biotin carbon atoms C-1 and C-7 would have the same labeling pattern. This is not the case (105, 182), and thus, the pimelate moiety must be assembled with one of the carboxyl groups covalently linked to another moiety. A thioester seems the most likely linkage (105, 182). Note that biotin is required for synthesis of malonyl-CoA, the postulated source of all of the carbon atoms of the pimelate moiety. Hence, we are presented with an evolutionary conundrum—biotin is required for biotin synthesis.
Biotin-requiring mutants of E. coli were first isolated many years ago. All of the mutants isolated as biotin auxotrophs were clustered at min 17 of the genetic map and defined five genes, called bioABCDF, based on mapping, cross-feeding, and complementation studies (47, 63, 178). However, during deletion analysis of the maltose utilization genes, a strain that required biotin was isolated and called bioH (184). More recently, strains having a nonfunctional pfs, the gene encoding 5'-methylthioadenosine/S-adenosylhomocysteine nucleosidase, were shown to require biotin due to accumulation of an inhibitory metabolite (see below) (34, 43). Biotin auxotrophs due to mutations in the bioH or pfs genes were not isolated by classical means probably because E. coli biotin auxotrophs require only miniscule amounts of this cofactor (supplementation with biotin at concentrations of a few nanomolars) and that conventional undefined growth media are often very rich in biotin. Therefore, several platings on biotin-free media are often required to detect the biotin requirement of auxotrophs. This raises the possibility that there are biotin synthetic genes that have yet to be identified.
Our knowledge of the early steps of biotin biosynthesis (Fig. 2) is in stark contrast to that of the late steps. The proteins (BioA, BioB, BioD, and BioF) of the late steps are well-studied enzymes, whereas the proteins of the early steps (BioC and BioH) are much more poorly understood. The sequence of steps in the late pathway was readily deduced since E. coli takes up each of the late intermediates. Growth of mutants with lesions in bioC and bioH proceeds when the medium is supplemented with 7-keto-8-amino pelargonic acid (KAPA) or any of the later intermediates in the pathway. No cross-feeding is observed between bioC and bioH mutant strains, suggesting that the early intermediates may not pass through cell membranes, perhaps because they are protein bound.
Essentially nothing is known of the function of BioC, a protein of 28.3 kDa. It is highly conserved among the proteobacteria and is often annotated as a SAM-dependent methyl transferase. However, since the 13C-labeling studies discussed above account for all of the carbon atoms of biotin coming from acetate and CO2 and were done in the presence of abundant methionine (thus precluding cycling of acetate to SAM), there is no straightforward role for a methyl transfer reaction in biotin synthesis. One possibility is that a methyl group is attached to a substrate or protein and is later removed. The BioC protein has not been studied biochemically probably because it readily forms inclusion bodies upon overexpression (200). It has been proposed that BioC acts as a carrier protein that carries an intermediate transferred by BioH (126), but there is no evidence for or against this notion.
In contrast to BioC, BioH is a well-behaved 28.5-kDa protein that allowed determination of its crystal structure at 1.7 Å (180). BioH is a monomeric two-domain protein (180, 200). A putative catalytic triad (Ser-82, His-235, and Asp-207) similar to that of the catalytic triad of hydrolases was identified. Moreover, in the BioH crystal structure the serine residue was found to have been modified by a protease inhibitor. Consistent with these indications of hydrolase activity, BioH had weak esterase activity on several model substrates (180), although this activity was not shown to depend on the Ser, His, Asp triad. Others had noted two Gly-Xaa-Ser-Xaa-Gly motifs in BioH that are characteristic of acyltransferase and thioesterase proteins (126). Thus, BioH seems likely to serve as an acyltransferase that may act as a hydrolase when water is the only available acceptor. However, the crystal structure gave no clues as to the identities of the substrates of BioH. BioH has been reported to bind CoA in vitro (200), but the significance of this finding is unclear. It should be noted that in the E. coli genome the bioH gene is well removed from the other genes of the pathway and is not regulated by the BirA repressor (see below). However, in other proteobacteria (e.g., the pseudomonads) bioH is found in a cluster of biotin synthetic genes that appear to be a transcription unit.
BioF is KAPA synthase, a pyridoxal phosphate-dependent homodimer of 41.6 kDa of known crystal structure (4, 217). The enzyme catalyzes the condensation of alanine with pimeloyl-CoA to give KAPA plus CoA and CO2 (resulting from decarboxylation of alanine). BioF is a two-domain protein with the pyridoxal phosphate bound in a crevice between the two domains formed by residues of both domains. The mechanism of the enzyme has been studied in some detail (216). Historically the enzyme has been assayed using pimeloyl-CoA, although pimeloyl-ACP could be the physiological substrate in E. coli (ACP-requiring enzymes will often use the analogous CoA compound as a model substrate). Consistent with this notion the E. coli KAPA synthase has a much higher Michaelis constant for pimeloyl-CoA than the analogous enzyme from Bacillus sphaericus (160), an organism in which pimeloyl-CoA is thought to be the physiological substrate due to the presence of pimeloyl-CoA synthetase.
BioA is 7,8-diaminopelargonic acid (DAPA) aminotransferase that has many similarities to BioF, the preceding enzyme in the pathway. Although the BioA subunit (47.3 kDa) is slightly larger than that of BioF, it is also a homodimeric pyridoxal phosphate-dependent enzyme. Indeed, the overall structure of BioA is very similar to that of BioF (116) and this is reflected in a weak sequence homology. BioA is a transaminase that converts KAPA to DAPA and as such is not a particularly interesting enzyme (193, 194). However, the amino donor is not a standard amino acid, but rather the highly activated amino acid S-adenosyl-2-oxo-4-thiomethylbutryate (SAM) (65, 194), which requires three ATP equivalents for its synthesis. The deaminated product derived from SAM spontaneously degrades in vitro (194), and thus, it seems likely that three ATP equivalents are consumed in what is an otherwise simple transamination reaction. The expense of this perplexing choice of amino donor may provide a rationale for the known tight regulation of biotin synthesis. However, it could be argued that use of a more pedestrian amino donor (B. subtilis uses lysine [210]) could alleviate the need for tight regulation.
In contrast to the preceding enzymes, BioD (dethiobiotin synthase or DTBS) catalyzes an unusually interesting step, the formation of the ureido moiety of biotin (70, 122). The BioD reaction is the ATP-dependent formation of dethiobiotin from DAPA and CO2. The enzyme is a homodimeric protein (subunit of 24.1 kDa) that is structured into a single well-folded domain (5, 101, 102, 179). X-ray crystallographic studies have shown that the reaction proceeds by carbamoylation of N-7 of DAPA (5, 101) (Fig. 3). Independent NMR evidence for carbamate formation has also been obtained (80). The second partial reaction is also unusual. In this reaction the carbamate is activated by transfer of the γ-phosphoryl moiety of ATP to a carbamate oxygen to form a mixed anhydride (Fig. 3). This mixed anhydride species has been demonstrated by time-resolved crystallography (115). The final step of the dethiobiotin synthase reaction is a nucleophilic attack by the N-8 nitrogen of DAPA on a carbamoyl oxygen of the mixed anhydride (Fig. 3), which results in formation of the ureido ring of dethiobiotin and release of the phosphate group.
The bête noire of biotin synthesis has long been the last step, insertion of the sulfur atom into DTB to form the thiophane ring of biotin. For many years this activity was ascribed to BioB by genetic means (biotin auxotrophs unable to grow on DTB) and could be assayed only by the ability of intact cells to convert DTB to biotin. Extensive attempts to obtain sulfur insertion in vitro all failed until Ifuku and coworkers (103) succeeded in showing biotin synthesis from DTB in a cell-free extract. The reaction required DTB, SAM, NADPH, BioB, and an unknown protein or proteins later shown to be flavodoxin (FldA) and flavodoxin reductase (Fpr) (24, 103, 104). This breakthrough was soon followed by demonstration of activity in a defined system containing NADPH, flavodoxin, and flavodoxin reductase as the electron transfer system plus DTB, SAM, and a BioB preparation as well as a reducing environment (181). BioB (a homodimer of a 38.6-kDa subunit) was found to be a very labile protein that is best purified and assayed under anaerobic conditions. The discovery that SAM was absolutely required for biotin synthesis and was not the sulfur donor (64) strongly suggested that BioB was a member of the (then) small family of "radical SAM" enzymes. It has recently become apparent that this is a fairly large family of proteins that catalyze a range of reactions that invariably involve difficult reactions accessible only by radical chemistry. The radical is generated by reductive cleavage of SAM to give a deoxyadenosyl radical (DOA˙) plus methionine. The DOA˙ radical then cleaves a C–H bond ?to generate a carbon radical that allows the chemistry to proceed. The electron donor in the single-electron reduction of SAM is a [4Fe-4S] cluster liganded to the cysteine residues of a perfectly conserved CX3CX2C motif. Consistent with this picture, the BioB reaction is chemically difficult since it requires cleavage and sulfur insertion into two unreactive C–H bonds. The mechanism currently accepted by most workers in the field (23, 108, 131) is given in Fig. 4, but it should be noted that the composition of BioB and the mechanism of its reaction are the subjects of a large and often conflicting literature.
The BioB species involved in the mechanism of Fig. 4 contains two [Fe-S] clusters. The number and composition of these clusters has been the subject of much disagreement in the literature. However, a variety of spectroscopic techniques plus a recent BioB crystal structure give a consistent picture. BioB contains two different clusters, the [4Fe-4S] cluster characteristic of radical SAM enzymes and a [2Fe-2S] cluster located at a different site. (The [2Fe-2S] cluster was often thought to be a [4Fe-4S] cluster degradation product until mutagenesis experiments suggested otherwise.) The crystal structure shows that BioB is an α/β8 (TIM) barrel protein with the two [Fe-S] clusters located at either end of the barrel (23). The [4Fe-4S] cluster is located at the open end of the barrel, whereas the [2Fe-2S] cluster (which utilizes an unusual arginine ligand) is at the closed end of the barrel. The crystal structures contain both SAM and DTB. The SAM is positioned such that reductive cleavage by the [4Fe-4S] cluster could readily occur while the DTB is positioned such that the C-9 carbon can accept a sulfur atom from the [2Fe-2S] cluster (23). This latter finding fits with the belief of most workers in the field that the [2Fe-2S] cluster is the immediate source of the biotin sulfur atom. This belief is supported by experiments in which each of the sulfur-containing small molecules of the defined in vitro reaction mixture was labeled with 35S and incorporation of the isotope into biotin was measured (see reference 84 and references therein). No radioactive biotin was obtained. Isotopically labeled biotin was obtained only when BioB was labeled with 35S in vivo (81) or with 34S by reconstitution of the [Fe-S] clusters in vitro (203). A more recent report showed that BioB reconstituted with Se in place of S gave selenobiotin derived from the [2Fe-2Se] cluster (205). Spectroscopic studies indicate that the [2Fe-2S] cluster disappears concomitantly with sulfur insertion (106, 202). The finding that BioB itself is the sulfur source impinges on one of the few points of agreement in the literature, that the BioB reaction is not catalytic in vitro (107, 108, 131). That is, despite extensive efforts over the fifteen years since the in vitro activity was first demonstrated, synthesis of more than one molecule of biotin per BioB monomer has not been reported. Numerous and diverse justifications have been put forth for the observed lack of catalysis, including absence of an essential cofactor (120), intrinsically inactive enzyme preparations (48), and inhibition by reaction products (151), but no general agreement has emerged. The currently favored and most provocative explanation for the lack of catalysis is that given above, the [2Fe-2S] cluster of the protein donates the biotin sulfur atom and this donation inactivates BioB. In this view BioB would be a reactant or substrate rather than an enzyme and, in the absence of repair of the [2Fe-2S] center, the protein would be sacrificed. The scenario of protein sacrifice is not completely unreasonable because there is no need for E. coli biotin synthase to be an efficient catalyst. E. coli (like most other organisms) requires only minuscule quantities of biotin for growth. E. coli can grow with only 100 to 200 molecules of biotin per cell (51, 128) and thus sacrifice of a few hundred molecules of a medium protein would not be a major drain on cellular resources. However, recent work has shown that E. coli BioB is catalytic in vivo. Choi-Rhee and Cronan (42) reported a series of experiments addressing whether or not biotin synthase is capable of multiple turnovers in E. coli cultures. Such in vivo measurements are difficult because the endogenous expression level of biotin synthase is very low and biotin may be split between pools of free and protein bound cofactor. The first issue was overcome by overexpressing hexahistidine-tagged biotin synthase (His6-BioB) under control of an arabinose-inducible promoter. The second issue was overcome by massively overexpressing, under control of an IPTG-inducible T7 promoter, biotin ligase (BirA) and a truncated, hexahistidine-tagged form of the acetyl-CoA carboxylase biotinyl domain that can accept biotin but does not form an active enzyme complex. These investigators then used a combination of antipentahistidine antibodies, [35S]methionine labeling, and streptavidin to quantify the levels of each protein and of total biotinylated protein separated by denaturing and nondenaturing gel electrophoresis. The use of the two gel systems allowed the turnover number of BioB to be calculated in an unusually straightforward manner. The ratio of biotinylated domain to BioB monomer gives 20 to 60 equivalents of biotin produced per initial biotin synthase monomer (42). The 20 turnovers observed in 4 h in vivo gives a minimum turnover number essentially equivalent to the maximal rate for one turnover observed in vitro (208). However, the in vivo measurement was complicated by the unexpected finding that enzyme turnover renders the enzyme susceptible to proteolytic degradation (42). A 50 to 90% depletion of the level of His6-BioB was observed after incubation. This depletion was not observed in the absence of DTB or in the presence of biotin (42). Thus, the view that BioB is sacrificed in its reaction remains germane.
The observed degradation of BioB was proposed to result from collapse of the enzyme [2Fe-2S] center due to donation of a sulfur atom to DTB. The [2Fe-2S] center of BioB is located deep within the barrel of this α/β8 (TIM) protein (23), and thus it seems probable that a substantial unfolding of the protein would be required to allow rebuilding of the [2Fe-2S] cluster. Such unfolding would allow restoration of the [2Fe-2S] center, but at the cost of exposure of the protein to proteolytic attack while unfolded. Therefore, in this scenario, catalysis by a molecule of BioB would require the protein to run a gauntlet of proteolysis until restoration of normal folding (with concomitant resistance to proteolysis) by rebuilding of the [2Fe-2S] center expended in biotin synthesis (42). The turnover numbers observed may thus be viewed as the products of a stochastic process. If the [2Fe-2S] cluster of a BioB molecule is rebuilt before proteolysis occurs, that protein will perform another turnover. If not, the protein molecule perishes and must be resynthesized de novo. Hence, some BioB molecules would catalyze only one or a few turnovers in their lifetimes whereas others may complete >100 turnovers.
The biotin requirement of pfs mutant strains (34) is due to inhibition of BioB by the by-product of sulfur insertion, 5'-deoxyadenosine (43). The pfs gene encodes the 5'-methylthioadenosine/S-adenosylhomocysteine nucleosidase, which was also shown to cleave 5'-deoxyadenosine to adenine plus 5'-deoxyribose (43). Mutants lacking Pfs activity precisely mimic BioB mutants in that they grow on biotin, but not on DTB or DAPA, and excrete DTB (43).
It is disconcerting that our understanding of the early steps of the E. coli biotin synthetic pathway remains so rudimentary. The mutants that define this part of the pathway, bioC and bioH, are predicted to encode a SAM-dependent methyltransferase and an esterase/acyltransferase, respectively. How can these putative activities be accommodated into a model that explains the predicted activities of these proteins and the 13C labeling results? The 13C labeling results strongly support a fatty acid synthetic mechanism for the pimeloyl moiety (105, 182). A malonyl unit (as a CoA or ACP thioester) would act as primer and retain the nonesterified carboxyl group. This primer would then be extended by two decarboxylating Claisen condensations with malonyl-ACP to give pimeloyl-ACP (127). This scheme is reasonable because type III polyketide synthases are known that use this malonyl-primed mechanism to make dicarboxylic acids of odd carbon lengths in which one the two carboxyl groups is in thioester linkage (11, 206). However, in the type III polyketide synthase reactions the internal ketone groups are retained in the final products (thus called polyketides) rather than being reduced to methylene groups as required to make the pimeloyl moiety. In biotin synthesis the keto groups would presumably be reduced by the usual reductive/dehydratase enzymes of fatty acid synthesis. However, the proposed intermediates are not typical fatty acid biosynthetic intermediates since they would have an odd number of carbon atoms and would have a carboxyl group in place of the usual methyl group. Hence, these substrates may have to be "disguised" to be recognized by the fatty acid synthesis enzymes. The annotations of BioC and BioH suggest the possibility that BioC converts the free carboxyl group to its methyl ester. Methylation would both cancel the carboxyl group charge and provide a methyl carbon that could mimic the methyl of the normal acyl chains. This methylated species would then have properties (chain length, hydrophobicity) approximating those of the substrates normally accepted by the enzymes of fatty acid synthesis. Following completion of the pimelic acid moiety the methyl ester would then be cleaved by BioH to give pimeloyl-ACP. However, there may be undiscovered biotin synthetic genes that play roles in the early reactions (see above). The Keio collection of single-gene knockout mutants of E. coli (12) should allow any additional biotin synthetic genes to be uncovered.
The BioB reaction also requires much additional study. This is a very intricate enzyme that cannot be considered understood. Although most investigators in the field agree with the general scheme of Fig. 4, there remain many loose ends. For example, the sequence of events that follow destruction of the BioB [2Fe-2S] cluster thought to donate the biotin sulfur atom remains unclear (202). Marquet and coworkers (202) reported that cluster destruction is accompanied by biotin formation, whereas others (106) report that biotin formation is 10- to 1,000-fold slower than cluster destruction and is biphasic. Another example is that one group has reported that 5'-deoxyadenosine is an extremely potent inhibitor of the in vitro reaction (151), in agreement with in vivo results (see above), whereas others report that it has no effect (204). It therefore seems possible that there may be several steps in the formation of biotin by BioB and different steps may be rate limiting in different enzyme preparations. A more extreme case is the claim that BioB has an intrinsic pyridoxal phosphate-dependent cysteine desulfurase activity responsible for generating the sulfur atom of biotin that would enter DTB via a persulfide (151, 152). This claim is countered by the finding that no pyridoxal phosphate is visible in the BioB crystal structure (23), that other laboratories have been unable to demonstrate pyridoxal phosphate binding or cysteine desulfurase activity (49, 206), and that biotin synthesis from DTB proceeds normally in cultures of E. coli starved for pyridoxal (2a). It should be noted that although the BioB crystal structure is a major step forward, crystallization necessarily selects for a single-protein species. Thus, the crystallized form of BioB may not fully represent all of the active enzyme species. Moreover, the present structure is of only moderate resolution (3.4 Å). An unsolved difficulty with the stoichiometry given by the BioB structure is that it contains only a single SAM molecule and there is no room for a second molecule (23). Therefore, the enzyme seems equipped to form only a single C–S bond. However, there are also disagreements with prior measurements of the stoichiometries of substrate binding and by-product formation (151, 207). It seems possible that, after generation of one carbon radical, the SAM could be regenerated, as occurs in the reactions catalyzed by two other SAM radical enzymes, lysine 2,3-aminomutase and spore photoproduct lyase (74). However, unlike BioB, the reactions catalyzed by these enzymes involve no overall change in the oxidation state of the substrate. Therefore, it seems more likely that following synthesis of the first C–S bond the methionine and 5'-deoxyadenosine products are released in order that a second molecule of SAM can bind.
The question of how the BioB [2Fe-2S] center is built and restored remains an open question. Although BioB has recently been reported to accept a [4Fe-4S] center from two E. coli Fe-S center scaffold proteins, SufA and IscA, no [2Fe-2S] center was formed (153). It should be noted that the BioB [2Fe-2S] has a novel ligand, an arginine residue rather than the Cys or His residues commonly used as ligands (23). This unusual ligand implied a specificity for the guanidium ligand, but recent results indicated that substitution of Cys, Ala, His, or Met for the arginine residue failed to inactivate BioB (26). Moreover, prior mutagenesis experiments indicated that two of the three conserved [2Fe-2S] cluster cysteine residues must be removed before BioB activity is lost (99, 153). The plasticity of this cluster suggests that the usual sulfur insertion pathways (the Isc and Suf systems) may not apply and, thus far, this seems to be the case. Inclusion of IscS does not allow BioB to become catalytic in vitro (120). The [2Fe-2S] cluster cannot be assembled by the Suf system in vitro (153) and E. coli strains with null mutations of either the suf or isc operons are not biotin auxotrophs (J. Imlay, personal communication). Unfortunately, suf isc double mutants are inviable so the possibility that biotin is synthesized due to redundant functions of the two systems cannot be tested.
Expression of the E. coli biotin synthetic (bio) operon is controlled by a simple, yet remarkably sophisticated, regulatory system in which the rate of transcription of the operon responds not only to the supply of biotin, but also to the supply of proteins (called biotin acceptor proteins) that become modified by covalent attachment of biotin (Fig. 5) (13, 14, 17, 37, 50, 52, 53). This regulatory system is understood in considerable detail thanks to a combination of genetic, physiological, biochemical, and biophysical investigations. The biotin operon of E. coli and other enteric bacteria is a striking example of regulation in which the transcriptional regulatory protein (BirA) is also an enzyme, in this case the biotin-protein ligase that catalyzes the covalent attachment of the essential vitamin, biotin, to certain proteins involved in key metabolic carboxylation and decarboxylation reactions. Moreover, regulation of the E. coli biotin operon is probably the best understood example of transcriptional regulation by an enzyme unrelated to nucleic acid metabolism. Superficially, the system resembles the classical TrpR regulation of the E. coli tryptophan operon where the Trp repressor protein binds to the trpEDCBA operator only when complexed with the corepressor, tryptophan. However in bio operon regulation, the repressor is also the biotin-protein ligase and the corepressor is not biotin, but biotinoyl-5'-AMP (bio-AMP), the product of the first half-reaction of the ligase reaction. It is these novel features that give this regulatory system its unusually subtle properties. The bio operon is actually two transcriptional units (bioA and bioBFCD) controlled by a common operator.
Maximal rates of bio operon transcription (derepression) occur when the biotin supply is severely limited (e.g., biotin starvation of a bio auxotroph) (Fig. 5A) or when high levels of a biotin acceptor protein are present (Fig. 5B). Under these conditions any bio-AMP synthesized is rapidly consumed in biotinylation of the acceptor protein (apo AccB) and hence no significant levels of the BirA-bio-AMP complex accumulate. Hence, BirA remains largely monomeric so the bio operator is seldom occupied and transcription is maximal. Repression of bio operon transcription occurs when the supply of biotin is in excess of that needed to biotinylate apoAccB. Under these conditions apo-BCCP is fully biotinylated, the BirA:bio-AMP complex accumulates, followed by dimerization of the protein to form the repressor species. The dimers then bind to the bio operator and repress transcription from both promoters. The two derepression conditions act by a common mechanism in that both decrease the levels of the BirA:bio-AMP complex available to bind the bio operator (Fig. 5C). Hence, the degree of repression of bio operon transcription can be most simply viewed as an antagonism between retention of bio-AMP in the BirA active site versus consumption of the bio-AMP bound to BirA by transfer of the biotinyl moiety to unmodified acceptor proteins (52). The model of Beckett and coworkers (17) in which the unmodified acceptor protein binds monomeric BirA and thereby inhibits formation of BirA dimers, the species required for effective repression, provides a structural context for this antagonism. Because the rate of bio operon transcription is sensitive not only to the intracellular concentration of biotin, but also to the supply of the protein to which the biotin must be attached, the net result of accumulation of the unmodified protein is an increase in the rate of synthesis of the small molecule needed for the posttranslational modification. The evidence for this model is strong and is discussed below.
The evidence that the ligase and repressor are the same protein was firmly established by data from several laboratories. The key genetic observation was that of Campbell and coworkers who showed that E. coli mutants defective in intracellular retention of biotin (called birA) were allelic to mutants defective in repression of the bio operon (called bioR; the birA designation has been retained). Since biotin is retained in E. coli only as the protein-bound species, it followed that the birA gene encoded biotin-protein ligase activity and this was demonstrated (13, 14). Furthermore, these workers also showed that a partially purified BirA protein preparation protected a specific segment of bio operon DNA (Fig. 6B) from nuclease digestion. This DNA segment contained a region of degenerate dyad symmetry previously defined as the operator of the bio operon (see below) by transcriptional (154) and mutational studies (16). As expected (see below), protection by BirA required the presence of bio-AMP. At about the same time, Eisenberg and coworkers showed that the purified repressor protein bound to bio operon DNA and catalyzed the biotin-protein ligase reaction (69). These workers also found that binding of the repressor protein to bio operon DNA in vitro required either biotin or bio-AMP but that bio-AMP was 1,000-fold more effective than biotin and biotin was active only at nonphysiological concentrations (161). Bio-AMP was also shown to be 1,000-fold more efficient than biotin in repression of bio operon transcription in a coupled transcription-translation system (162). Since these pioneering studies, it has become possible to obtain large amounts of BirA (normally a very nonabundant protein) (32, 229), which has led to biophysical studies as well as crystal structures of the unliganded (apo) protein (222) and of complexes of BirA with biotinoyl-lysine (222), biotin (215), or a nonhydrolyzable analogue of bio-AMP (227). Although we lack the structure of the tertiary complex of BirA, the bio operator and bio-AMP (or an analogue), these studies show that BirA is a winged helix-turn-helix protein (198, 222) of 35.2 kDa (Fig. 6). The winged helix-turn-helix is located at the extreme N terminus of the protein and is one of the three BirA domains, the others being a large central domain where an active site is found and a small C-terminal domain. The latter two domains show high levels of structural similarity with biotin-protein ligases from throughout biology (38). More recent work has shown that BirA requires bio-AMP to dimerize at physiological concentrations (71) and only the BirA dimer can efficiently bind the operator (1, 195, 197, 199). Bio-AMP binding activates the assembly of the BirA-operator complex by increasing the extent of dimerization by three orders of magnitude (124, 196).
The biotin attachment activity of BirA (Fig. 7) proceeds through the bio-AMP intermediate formed from biotin and ATP (38). Enzyme-bound bio-AMP is then attacked by the ε-amino group of a specific lysine of the acceptor protein to give the biotinylated acceptor protein (38) (Fig. 7). In the absence of an appropriate acceptor protein the bio-AMP intermediate remains bound within the BirA active site where it is quite stable (229). BirA shows very high specificity for biotin. The discrimination in favor of biotin versus DTB is ca. 50,000-fold (42, 228), although BirA-catalyzed attachment of DTB can be demonstrated (228). Both DTB and the oxidized form of biotin, biotin sulfoxide, show very weak abilities to derepress transcription of the biotin operon (35). A large number of birA mutants have been isolated based on their transcriptional phenotypes (using bio-lacZYA fusions) (15) and the mutational alterations of a considerable number of these have been determined by DNA sequencing (31). These fall into three main classes: mutants defective in regulation (the classical bioR phenotype), mutants defective in binding biotin and/or bio-AMP (the classical birA phenotype [36]), and those having temperature-sensitive growth (15). However, there is considerable overlap among these phenotypes and some mutant proteins show all three phenotypes (15). All BirA crystal structures including that with a bio-AMP analogue show the N-terminal DNA binding domain markedly protruding from the body of the protein (Fig. 6A) and thus it is surprising that deletion of this domain has a profound effect on the ligase activity of the truncated protein due to poor binding of biotin and/or bio-AMP (230). It should be noted that BirA is an essential gene (12, 15, 79) since it is required for fatty acid synthesis and, hence, membrane lipid synthesis (56).
AccB protein, the sole biotin acceptor protein of E. coli, is an unusual protein; the N-terminal half appears largely unstructured, although the extreme N terminus is known to interact with the AccC subunit (41), whereas the C-terminal half of the protein is folded into a compact and stable structure called the biotin domain (Fig. 7). This domain has a structure similar to that of lipoyl domains (see below). The AccB biotinoyl domain is as efficient a biotin acceptor as the full-length protein (144) and is often used for in vitro work to avoid the problems with aggregation of the full-length protein (41, 144). The structure of biotinoyl domains is strongly conserved throughout biology, and expression of foreign biotinoyl domains in E. coli can derepress bio operon transcription (55). Mutants of the AccB biotinoyl domain have been isolated that are defective in interaction with BirA (39), and mutations have been introduced that allow the protein to accept lipoic acid in place of biotin (168). The work on the stucture and function of the biotin domain is intimately involved with (and is historically derived from) that on lipoyl domain structure and will be discussed in that context below.
The enzymes of E. coli biotin synthesis are encoded (with the exception of bioH) by a cluster of genes located adjacent to the attachment site of phage λ called the biotin (bio) biosynthetic operon (Fig. 5 and 6). Transcription of these bio genes is from two partially overlapping face-to-face promoters controlled by a common operator site of 40 bp that binds a dimer of the BirA protein (16, 17, 130, 154) (Fig. 6B). The leftward promoter transcribes bioA, whereas the rightward promoter transcribes bioBFCD. The 5' ends of the transcripts have been mapped, and mutations within the operator that ameliorate repression of either rightward or leftward transcription (or both) are known (16, 118, 154, 186). The operon and operator sequence are conserved in S. enterica and Citrobacter freundii (186). A long-standing puzzle is that bioH is not under BirA regulation (15), especially since in other proteobacteria (e.g., pseudomonads) bioH seems part of a biotin biosynthetic operon. Also, unlike many repressors, BirA does not appear to be autoregulated because it is cotranscribed with an essential gene (murB) involved in peptidoglycan biosynthesis.
E. coli contains only a single species of biotin acceptor protein, the AccB subunit of acetyl-CoA carboxylase (ACC), which is the first enzyme of fatty acid biosynthesis (50, 51, 54, 72) and is therefore essential for growth. The response of the E. coli biotin regulatory system to the supply of biotin acceptor proteins is readily rationalized by the fact that biotin attains biological function only when the vitamin is covalently attached to AccB; the free vitamin cannot support ACC activity (56). AccB, which is also called biotin carboxyl carrier protein (BCCP), forms an unstable complex with AccC, the subunit that catalyzes the biotin carboxylase partial reaction of acetyl-CoA carboxylase. The chromosomal locations of the genes (accA and accD) that encode the other two ACC subunits are well removed from the accBC operon and each other (50, 51, 54, 72). The AccB-AccC complex was recently shown to consist of an AccC dimer plus four copies of AccB (41). This complex is thought to bind an α2β2 heterotetramer of the AccA and AccD subunits to give active ACC, the enzyme required for production of malonyl-CoA, the key precursor of fatty acid synthesis (56). The rates of transcription of all four genes are controlled by cellular growth rate (129), which is physiologically reasonable since lipids (hence, fatty acids) constitute a constant fraction of the cell mass. The fact that bio operon transcription is derepressed by increased synthesis of AccB nicely ties biotin synthesis to growth rate. This is because increased growth rates require increased flux through the fatty acid synthetic pathway in which ACC catalyzes a rate-limiting step (62). Indeed, biotin consumed by increased protein biotinylation has been shown to be restored by increased biotin synthesis (53).
The fact that the only acc genes that are cotranscribed are accB and accC and that this gene arrangement is very widely conserved in bacteria raised the question of its relevance to the regulation of biotin synthesis (2). It seems possible that the defined stoichiometry given by cotranscription of accB and accC might function to aid efficient biotinylation of AccB. It seemed possible that an excess of AccC might tie up apo-AccB in a complex that would be a poor substrate for BirA and thereby disrupt the regulatory system (Fig. 5D). This has been shown to be the case (2). Overproduction of AccC gave almost maximal repression at biotin concentrations that normally give only slight repression and inhibited biotinylation of AccB. As expected overproduction of both AccB and AccC to restore the normal ratio of the two proteins relieved the down-regulation given by overproduction of AccC alone and this relief required that the overproduced AccB species be competent to interact with AccC (2).
The present model of bio operon regulation has a very solid experimental basis obtained by both in vivo and in vitro approaches. However, there are two views of the mechanism whereby accumulation of the unmodified biotin domain derepresses transcription of the operon. In one view this is simply a competition for bio-AMP between its consumption by protein biotinylation versus its presence in the BirA active site where it triggers dimerization and subsequent operator binding (52). In the second view the biotin domain forms a heterodimeric complex with a monomer of BirA. The BirA surface used to form the heterodimer is proposed to be the same surface as that used in forming the BirA homodimer. Hence, in this view, competing protein-protein interactions are responsible for derepression on accumulation of unmodified biotin domain (17). The conceptual distinction between these two models is the lifetime of the BirA-biotin domain interaction. In the bio-AMP competition model the interaction is ephemeral, the two proteins associate, biotin is then transferred, and the complex rapidly dissociates, whereas in the competing protein-protein interaction model the BirA-biotin domain interaction is long lived. One approach that might distinguish these models would be to use the small peptides that are substrates for biotinylation by BirA (183). These peptides, which were isolated by screening large peptide libraries, are quite diverse in sequence and have as few as two residues (one being the reactive lysine residue) that are found in naturally biotinylated proteins (183). Due to their small sizes (14 residues is sufficient [18, 183]) and diverse sequences, it seems unlikely that stable peptide-BirA complexes are made. If these sequences (attached to a partner protein) are expressed in E. coli they should derepress bio operon expression if the bio-AMP competition model is to be supported. If they fail to derepress, but are efficiently biotinylated, then the competing protein-protein interaction model would be supported. Although the most studied of these peptides is reported to be as good a biotin acceptor as the AccB biotin domain, this peptide remains enigmatic because it seems to lack structure in solution (18) and can only be biotinylated by BirA (55). Biotin ligases from other organisms fail to use this peptide as a biotin acceptor, although these ligases readily utilize the AccB domain as a substrate (55).
Another approach to further our understanding of bio operon regulation would be studies of a missing class of BirA mutants, BirA superrepressing mutants. These would be mutants that would repress transcription under all conditions, including biotin limitation and apo-domain overexpression. Some of the possible classes of mutants are: (i) BirA proteins unable to bind the biotin acceptor protein, (ii) BirA proteins that bind the acceptor protein but are unable to biotinylate it, (iii) BirA proteins that form very tight homodimers (perhaps even in the absence of bio-AMP), and (iv) BirA proteins that cannot dissociate from the operator DNA. Some of the mutants might be genetically dominant. Most of these mutants would be nonviable because fatty acid synthesis would be blocked due to lack of biotinylation of AccB, which would account for the fact that such mutants have not been reported. Hence, the isolation of superrepressor mutants would require expression of a heterologous biotin protein ligase active on AccB to allow fatty acid synthesis to proceed. Depending on the types of mutants isolated these could significantly refine our knowledge of the bio regulatory system.
Finally, the crystal structure of BirA complexed with the bio operator and bio-AMP (or an analogue) seems likely to be very informative. This may give information on the conformational changes in BirA that accompany bio-AMP binding (29). Cocrystals of the BirA-biotinoyl domain complex would also be of great interest.
Lipoic acid (Fig. 1) is a sulfur-containing cofactor found in most prokaryotic and eukaryotic organisms. In E. coli and other organisms lipoic acid is essential for function of several key enzymes involved in oxidative and single-carbon metabolism, including pyruvate dehydrogenase (PDH), 2-oxoglutarate dehydrogenase (2-OGDH), branched-chain 2-oxoacid dehydrogenase, acetoin dehydrogenase, and the glycine cleavage system (173). In each enzyme, a specific subunit is modified by attachment of lipoic acid to specific lysine residues within conserved domains of these subunits. In each of these domains an amide linkage is formed between the carboxyl group of lipoic acid and the ε–amino group of the specific lysine residue (121). During catalysis, the protein-bound lipoamide moieties serve as carriers of reaction intermediates among the multiple active sites of these multienzyme complexes (173).
Our knowledge of the pathways of lipoic acid synthesis, attachment, and function has progressed rapidly in the past 10 years largely due to complementary genetic and biochemical analyses in E. coli. I will first discuss the enzymes that carry and require the cofactor because they are derived from diverse areas of metabolism. Next, the mechanisms of attachment of lipoic acid and its precursor, octanoic acid, to these proteins will be reviewed. Finally, the synthesis of the cofactor itself will be discussed. This organization was chosen because the unusual biosynthetic pathway of lipoic acid is mechanistically intertwined with attachment of the cofactor.
The PDH reaction mechanism is probably the most thoroughly characterized lipoic acid-dependent enzyme. PDH catalyzes the oxidative decarboxylation of pyruvate to the key metabolic intermediate, acetyl-CoA. This very large enzyme complex consists of multiple copies of each of three subunits encoded by the aceE aceF lpd operon. The first subunit (AceE) is a thiamine diphosphate-dependent decarboxylase (E1p) that catalyzes both the decarboxylation of pyruvate and the reductive acetylation of the lipoyl group that is covalently attached to the second subunit, E2p (AceF). The E2p subunit is a dihydrolipoyl acetyltransferase responsible for the transfer of the acyl group from lipoyl moiety to CoA to form acetyl-CoA. The third subunit, E3 (Lpd), is a dihydrolipoyl dehydrogenase that serves to regenerate the disulfide bond of the lipoyl moiety of E2p (158) and thereby prepares the enzyme for another cycle of catalysis. The E2p subunit to which E1 and E3 are bound strongly (but noncovalently) forms the structural core of the multienzyme complex. The oxidative decarboxylation of pyruvate to form acetyl-CoA is the link between glycolysis and the citric acid cycle, and therefore, PDH activity is essential to cells that rely on respiration to provide metabolic energy. In most aerobically respiring organisms the PDH complex also supplies the acetyl-CoA necessary to sustain essential biosynthetic pathways, especially those of fatty acid and amino acid synthesis (90). Synthesis of the PDH complex varies over a 7- to 10-fold range depending on the growth conditions (66, 125, 187). It is induced by exogenous pyruvate or when pyruvate is generated endogenously, e.g., by thiamine starvation, and it is partially repressed by excess glucose and during growth on acetate or on citric acid cycle intermediates. Regulation by pyruvate or a derivative of pyruvate proceeds through the PdhR repressor (164, 165). PDH synthesis is repressed during anaerobic fermentative growth where the catalytic activity is also inhibited. Under these conditions the conversion of pyruvate to acetyl-CoA is mediated by the derepression and activation of pyruvate formate lyase (86, 187).
The mechanism of 2-OGDH is essentially the same as that of PDH, as is the structure of the complex. Indeed, the 2-OGDH complex has been reported to contain low levels of PDH subunits (190). The 2-OGDH complex contains three subunits: a 2-oxoglutarate decarboxylase component (E1o), a trans-succinylase component (E2o), and a dihydrolipoyl dehydrogenase (E3). The E1o and E2o subunits are different proteins from the corresponding subunits of the PDH complex and are encoded by the sucA and sucB genes, respectively. However, the E3 subunit is the same protein, Lpd, found in the PDH complex. In aerobically grown E. coli, this complex catalyzes a key step in the citric acid cycle and also supplies succinyl-CoA for biosynthesis of two amino acids, methionine and lysine (97). Under the appropriate conditions, E. coli strains lacking functional 2-OGDH can be supplemented with succinate or methionine plus lysine to provide metabolic bypasses of loss of this enzyme complex (97). Expression of the 2-OGDH is highly induced during aerobic growth on acetate and citric acid cycle intermediates and is severely repressed during fermentative growth where succinyl-CoA is generated by succinyl-CoA synthetase (90), although 2-OGDH is synthesized in anaerobic media containing an electron acceptor such as nitrate or fumarate (163).
The third lipoylated protein of E. coli is the H protein of the glycine cleavage system, an enzyme widely distributed in bacteria and in the mitochondria of plants (where it is called glycine decarboxylase), fungi, and mammals (67, 76, 148). The glycine cleavage system catalyzes the reversible cleavage of glycine, yielding carbon dioxide, ammonia, 5,10-methylenetetrahydrofolate, plus a reduced pyridine nucleotide. It consists of four component proteins termed the T, H, P, and L proteins. The first three proteins are encoded by the gcvT gcvH gcvP operon, while L protein is the same as Lpd, the E3 protein of the 2-oxoacid dehydrogenases as discussed above (191). P protein catalyzes the pyridoxal phosphate-dependent decarboxylation of glycine and transfers the remaining methylamine moiety to one of the sulfhydryl groups of the lipoyl prosthetic group of H protein. T protein catalyzes the release of ammoniate and transfer of the one-carbon unit to tetrahydrofolate from the lipoyl residue. L protein is a lipoamide dehydrogenase that catalyzes the reoxidation of the dihydrolipoyl residue of H protein and reduction of NAD+. Thus, the lipoic acid moiety of H protein interacts with the active sites of three different enzymes in a manner analogous to that found for 2-oxoacid dehydrogenase complexes.
In all 2-oxoacid dehydrogenase complexes, the core of the structure is provided by the E2 subunit to which the E1 and E3 components are bound tightly but noncovalently. In the PDH and 2-OGDH complexes of E. coli and other gram-negative bacteria (57, 93) plus the 2-OGDH and branched-chain 2-oxoacid dehydrogenase complexes of mammals (85, 92), the core consists of 24 copies of the E2 chain arranged with octahedral symmetry, whereas in the PDH complexes of mammals and gram-positive bacteria (8, 96, 117, 132), the core comprises 60 E2 chains arranged with icosahedral symmetry. In all 2-oxoacid dehydrogenase complexes, the E2 component has a multidomain structure comprising (from the N terminus): a lipoyl domain (or domains of ca. 9 kDa), a small peripheral subunit-binding domain (ca. 4 kDa), and a much larger catalytic domain (ca. 28 kDa) that houses the acyltransferase activity and aggregates to form the inner core of the complexes. These domains are separated by long (25- to 30-residue) segments of polypeptide chain, characteristically rich in alanine, proline, and charged amino acids that form flexible but extended linkers (158).
The numbers of PDH lipoyl domains per E2 subunit varies from one to three. In the PDH complexes of gram-negative bacteria, the number is usually three (e.g., E. coli and Azotobacter vinelandii) or two (e.g. Haemophilus influenzae, Neisseria meningitidis, Alcaligenes eutrophus, and Thiobacillus ferrooxidans) (159). All of the 2-OGDH E2o subunits described to date contain a single lipoyl domain, as is also the case for the E2b chains of all BCDH complexes (19, 158, 159, 173). A generally applicable explanation for the variation in the number of lipoyl domains has not yet been worked out. Protein-engineering experiments have eliminated the straightforward explanations. In E. coli PDH, selective deletion of one or two lipoyl domains has no detectable effect on the overall catalytic activity, the system of active-site coupling, or the ability to complement pyruvate dehydrogenase complex mutants (89). As expected, the catalytic activity is abolished when all three lipoyl domains are deleted or when the lipoyl domains are rendered unlipoylatable by conversion of the lipoylated lysine residue to glutamine (9, 89). There is no mandatory order of reductive acetylation of the repeated lipoyl domains within E2p polypeptide chains because complexes containing mixtures of wild-type and mutant lipoyl domains (+/−; −/+; +/+/−) are fully active, although the complex containing the −/−/+ version of the E2p polypeptide chain showed a 50% reduction in specific activity (9). Activity is also impaired (but not abolished) by increasing the lipoyl domain content to four to nine per E2p chain, possibly due to underlipoylation of the domains participating in catalysis and interference from unlipoylated domains (135). High-field NMR studies were carried out with variants containing zero to nine lipoyl domains per E2p subunit. These studies suggest an explanation for the presence of three lipoyl domains per E2p subunit in the wild-type PDH complex that is based on the greater inherent mobility and thus potentially more efficient active-site coupling of this arrangement (136). The superiority of the three lipoyl domain-PDH complex has since been confirmed by physiological studies, from which it was concluded that decreased lipoyl domain contents adversely affect growth rate and growth yield (61). The physiological consequences of increasing the number of lipoyl domains from three to nine per E2p chain, and the effects of inserting up to seven unlipoylated (mutant) domains between a wild-type N-terminal lipoyl domain and the E3 binding domain, were also investigated, and the findings indicate that three lipoyl domains per E2p chain are optimal and that only the outermost lipoyl domain needs to be lipoylated to obtain full catalytic activity (87). It was concluded that the reason for retention of three lipoyl domains is to extend the reach of the outermost lipoyl cofactor rather than to provide extra cofactors for catalysis (87). However, given this advantage, why then do many lipoylated proteins contain only a single lipoyl domain?
The conserved structure of lipoyl domains (Fig. 8A) is directly related to catalytic functions of the domain in substrate channeling and active-site coupling. First, although free lipoic acid is a substrate for E2p and E3 in vitro, lipoylated domain is a much better substrate (82). Attachment of the lipoyl group to the conserved lysine at the tip of the protruding β-turn gives a dramatic reach to the "business end." Moreover, the flexible and extended linker regions that connect the lipoyl domain(s) with the catalytic domain contribute increased mobility to the swinging arm, since deletion of the linker region in a modified "single lipoyl domain" E2p chain caused an almost total loss of overall activity without substantially affecting the individual enzymatic activities (140). Second, E1p and E1o of E. coli (85, 114) and A. vinelandii (22) can only transfer acyl groups to their cognate E2 protein, thereby providing an accurate substrate-channeling mechanism such that the reductive acylation occurs only on the lipoyl group covalently attached to the appropriate E2 subunit. Third, although the attached lipoate was once thought to be freely rotating (59, 82), recent structural data showed that the lipoyl-lysine β-turn of the domain became less flexible after lipoylation of the lysine residue (110). The restricted motion of the lipoyl group would facilitate the effective E1 and E2 interaction by presenting the lipoyl group in the preferred orientation to the active sites of E1 and thereby enhance catalysis. This is in agreement with the recent crystal structure of the E1 component of the branched-chain acid dehydrogenase complex from Pseudomonas putida (3). According to this structure, the active site where thiamine diphosphate binds is at the bottom of a long funnel-shaped tunnel, which suggests that the lipoyl group attached to the lipoyl domain must be fully extended and accurately positioned to reach the thiamine diphosphate cofactor. Amino acid side chains responsible for this specific positioning have been mapped to two residues that flank the lipoyl-lysine (214). Finally, the prominent surface loop connecting β-strands 1 and 2 (which lie close in space to the lipoyl-lysine) is another major determinant of the interactions of the lipoyl domain with E1 (213). Deletion of this loop results in a partially folded domain and almost completely abolishes lipoylation and reductive acylation, indicating that the loop is involved in maintaining the structural integrity of the domain, posttranslational modification, and catalytic function (110). It is believed that the loop structure is important for stabilizing the thioester bond of the acyl-lipoyl intermediate (109, 110).
Subgenes that encode the lipoyl domains from a wide range of bacteria, including E. coli E2p (6) and E2o (176), Bacillus stearothermophilus E2p (60), human E2p (166), A. vinelandii E2p (19) and E2o (21), and N. meningitidis E2p (201), have been overexpressed in E. coli, and sufficient recombinant protein has been obtained for the domain structures to be determined by multidimensional nuclear magnetic resonance (NMR) spectroscopy. The archetypical structure, that of the single apo-lipoyl domain of the E2p chain of B. stearothermophilus (58), is composed largely of two four-stranded β-sheets, with the N- and C-terminal residues of the domain close together in space in one sheet and the lysine residue earmarked for lipoylation in an exposed position in a tight type I β-turn generated by β-strands 4 and 5 in the other sheet. There is a well-defined hydrophobic core, the least well-defined regions being the exposed β-turn where the lipoyl-lysine resides and, most notably, the nearby large surface loop that connects β-strands 1 and 2 (Fig. 8A). Consistent with the high level of sequence similarity between lipoyl domains of 2-oxoacid dehydrogenase multienzyme complexes, all other lipoyl domains conform to the same structural pattern. Given the small differences in the NMR spectra of the lipoylated and unlipoylated forms of the B. stearothermophilus (59) and A. vinelandii (20) E2p domains, the structures of holo- and apo-domains have been inferred to be substantially the same.
The determination of lipoyl domain structures has allowed prediction of the structure of another lipoylated protein: the H protein of the glycine cleavage system. H proteins are about 130 resides in length (75). Although the overall sequence identity was low (<20%) (77), the conservation of key residues indicated that there was likely to be considerable structural similarity between the H protein of the glycine cleavage system and the lipoyl domains of 2-oxo acid dehydrogenase complexes (27). Indeed, the X-ray crystal structure of the lipoylated pea leaf H protein agreed well with the theoretical predictions. The biotinyl domains of biotin-dependent enzymes have structures strikingly similar to those of lipoyl domains (Fig. 8B) as originally predicted by Brocklehurst and Perham (27). This is particularity true of biotin domains from enzymes other than bacterial and plant plastid acetyl-CoA carboxylases. The biotinylated subunits of the bacterial and plastid acetyl-CoA carboxylase contain a characteristic thumb structure not found in other biotinoyl domains or in lipoyl domains (51). The structure of the biotin domain of E. coli AccB has been established by X-ray crystallography (10) and NMR spectroscopy (Fig. 8B) (177, 231, 232). The structure closely resembles those of the lipoyl domain in the E2 component of 2-oxoacid dehydrogenase complexes and of the H protein in the glycine cleavage system. Like these lipoylated proteins, the AccB domain is a flattened β-barrel, comprising two four-stranded antiparallel β-sheets, with the biotinyl-lysine residue located in the exposed β-turn between β-strands 4 and 5 (Fig. 8B). The high-resolution NMR structure of another biotinoyl domain, that of Propionibacterium shermanii transcarboxylase, has also been determined (170). This structure more closely resembles the lipoyl domain structures since it lacks the protruding thumb of the E. coli biotin domain (to which it is otherwise quite similar). Depending on the pair of domains chosen for comparison the root-mean-square deviation of biotinoyl and lipoyl domain backbone atoms can be as low as 1 Å; hence, these proteins define a protein family (PF00364). Other work has shown that one of the proline/alanine-rich linker regions that lies between the domains of E. coli PDH can functionally replace the proline/alanine-rich linker region that lies upstream of the biotin domain of E. coli BCCP (54), underlining the interrelatedness of the biotin and lipoic acid acceptor proteins.
Posttranslational modification of apoproteins with lipoic acid occurs by several mechanisms. In E. coli, two complementary systems for protein lipoylation have been characterized by genetic and subsequent biochemical analyses. Exogenous lipoate or octanoate is transferred to unlipoylated apoproteins in an ATP-dependent process by lipoate-protein ligase (LplA) (142, 143). The second E. coli pathway requires the lipB gene product (octanoyl-ACP:protein-N-octanoyltransferase) to transfer endogenously synthesized octanoate to apoproteins, which then becomes the substrate for sulfur insertion (Fig. 6) (45, 46, 113, 142, 234).
Lipoate-protein ligase activity was first described by Reed and coworkers (175) in Enterococcus faecalis, as well as in E. coli, and these workers postulated that lipoate was attached to protein by a two-step ATP-dependent reaction with lipoyl-AMP as an activated intermediate. The reaction is exactly the same as that of BirA (Fig. 7) with the substitution of lipoic acid for biotin (hence, this is not shown). Although the lipoate-protein ligases were key reagents in demonstration of the role of lipoic acid in the 2-oxoacid dehydrogenase reactions (121, 174), neither protein had been purified to homogeneity and thus the proposed mechanism could not be proved. The E. coli lplA gene was the first lipoate-protein ligase gene to be isolated, and LplA was the first such ligase purified to homogeneity (83, 142). lplA mutants were isolated by two different approaches. In the first approach a lipA strain was mutagenized by transposon insertion and the mutagenized cells were supplemented with a mixture of succinate and acetate to bypass the lipoate requirement. The supplement was then switched to lipoate, and an ampicillin enrichment was performed followed by plating onto the succinate-acetate-supplemented medium. The resulting colonies were screened for strains able to grow on succinate-acetate-supplemented medium but not on lipoate-supplemented medium. Three classes of such mutant strains could have resulted from this scheme: strains lacking the ligase (lplA), strains defective in lipoate transport, and lpd mutants that lack the E3 subunit common to all of the lipoate-dependent enzymes of E. coli. Indeed, the selection was an unwitting repeat of the selection used for lpd mutants (88). Surprisingly, all of the mutants isolated were lplA mutants. It is unclear why no lpd mutants were isolated in the lplA selection and vice versa. The lack of lipoate transport mutants suggests that there may be no lipoate transporter in E. coli (as is believed to be the case for short-chain fatty acids). Given the small size, the hydrophobicity, and the miniscule amount of the cofactor needed for growth, no transporter may be needed. Indeed, it has been reported that both enantiomers of 35S-lipoate were taken up by E. coli, although only R-lipoic acid became attached to the 2-oxoacid dehydrogenases (147). Since a transporter protein would be expected to discriminate between enantiomers, this finding argues strongly against the existence of a lipoate transporter. Mutants mapping in lplA were also isolated by a direct selection, resistance to selenolipoic acid. Selenolipoic acid is a growth-inhibitory lipoate analogue in which the sulfur atoms are replaced with selenium (172). These mutants proved to encode a ligase of somewhat compromised activity that was able to discriminate against the selenium analogue (143).
The purified LplA enzyme is a 38-kDa monomeric protein (83). Assays with a fully purified apoprotein acceptor have demonstrated that purified LplA plus lipoate and Mg-ATP are sufficient to reconstitute lipoylation in vitro (83, 142, 168, 169). Thus, it is clear that LplA catalyzes both the ATP-dependent activation of lipoate to lipoyl-AMP and the transfer of this activated lipoyl species to apoprotein with concomitant release of AMP. This conclusion is consistent with the early findings of Reed and coworkers (175) that the E. coli enzyme could not be fractionated into separable lipoate activation and lipoyl transferase components. The LplA enzyme has been shown to be capable of utilizing lipoate and several lipoate analogues as donors for the posttranslational modification of E2 apoproteins in vivo (142). This rather broad substrate specificity in vivo matches the similarly broad substrate specificity observed (28).
Very recently, ten crystal structures of LplA and of LplA homologues were reported, including structures of E. coli LplA (78) plus an LplA-lipoic acid complex (78). The reported structures agree well and show E. coli LplA to be a two-domain protein consisting of a large N-terminal domain and a small C-terminal domain. The E. coli LplA-lipoic acid complex was difficult to interpret because lipoic acid was bound to be different LplA molecules within the crystals by different modes and with poor resolution. For example, in one case, the lipoic acid carboxyl was hydrogen bonded to Ser-72, whereas in the other case Arg-140 was the hydrogen bond donor (78). Since enzymes rarely show such plasticity and lipoic acid is a hydrophobic molecule, it seemed possible that the observed association with a hydrophobic LplA surface in the interdomain cavity was artifactual. Moreover, in prior work, Reed and coworkers had isolated LplA mutants resistant to selenolipoic acid (172). Since this is a very discrete modification of the LplA substrate, the mutant protein would be expected to have an alteration close to the pocket that binds the lipoic acid thiolane ring. However, the site of this mutation (Gly-76 to serine [143]) was distal from the lipoate binding site reported. This dilemma appears resolved by two lipoic acid-containing structures of an LplA homologue from the archaeon Thermoplasma acidophilum (119, 139) that can be readily superimposed on the E. coli LplA structure, except that the T. acidophilum protein lacks the LplA C-terminal domain. In both T. acidophilum structures the lipoate thiolane ring was close to the glycine residue that corresponds to E. coli Gly-76, the residue giving resistance to the selenium analogue, and a plausible reorganization of the molecule to prevent binding of the larger selenolipoic acid was proposed (139). Moreover, addition of lipoic acid to a complex of the T. acidophilum with ATP gave lipoyl-AMP, thereby showing that the lipoic acid was bound in a physiologically meaningful manner (119). Lipoyl-AMP was bound in a U-shaped pocket and was well shielded from solvent. The T. acidophilum LplA was reported to be inactive in catalyzing the overall LplA reaction (139), although lipoyl-AMP synthesis was demonstrated (119). Since T. acidophilum LplA lacks the C-terminal domain of E. coli LplA (119, 139), this suggested that the missing domain plays a key role in transfer of the lipoyl moiety from lipoyl-AMP to the acceptor domain. Indeed, a second protein has been proposed to interact with T. acidophilum LplA and allow the complete reaction (139). The fact that the lipoate of one of the T. acidophilum LplA structures was converted to lipoyl-AMP and that the locations of lipoate moieties of the two T. acidophilum LplA structures agreed well argues that these represent the catalytically competent lipoate binding site. It therefore follows that the lipoate of the E. coli LplA structure bound in a different area of the LplA molecule is bound in a catalytically inappropriate manner. The structures of two LplA homologues from Streptococcus species have also been reported to the Protein Data Bank (PDB ID codes 1VQZ and 2P0L). Both proteins have structures very similar to that of the E. coli protein.
During the characterization of E. coli lplA null mutant strains, compelling evidence was found for a second protein lipoylation pathway that did not require the lplA gene product. When independently derived lplA null alleles were transduced into wild-type strains, the resulting mutant strains showed no growth defects on minimal glucose medium, indicating that these strains possessed functional (therefore lipoylated) 2-oxoacid dehydrogenases. This was directly confirmed by bioassays that showed lplA null mutants to contain normal levels of lipoylated proteins (143). Thus, it was clear that E. coli has an lplA-independent lipoylation pathway. This was first attributed to a second ligase that had somehow been missed in the biochemical analyses, perhaps due to the in vitro conditions chosen. However, no such second ligase could be found (83), and thus alternative pathways were considered. The most straightforward alternative pathway was that the fatty acid synthesis intermediate, octanoyl-ACP, would be converted either directly or indirectly to lipoylated proteins. That is, lipoate synthesis would occur without a free carboxyl group. The carboxyl group would be bound in the thioester bond that links fatty acids to ACP and this bond would then be attacked by the ε–amino group of the lipoyl domain lysine residue to give the amide linkage. Several lines of evidence demonstrated that this alternative protein lipoylation pathway depended on the lipB gene product. The lipB gene was originally isolated as a class of lipoic acid auxotrophs (211). These mutants showed residual (leaky) growth in the absence of lipoic acid despite having putative null mutations due to transposon insertions into lipB (171, 211). This leakiness was reflected in their 2-oxo acid dehydrogenase activities and lipoylated protein contents. These strains retain about 20% of the enzyme activities and about 10% of lipoyl protein content of wild-type strains (171). The leaky growth of lipB strains in the absence of lipoate was eliminated by introduction of an lplA mutation, suggesting that lipB was involved in lipoyl domain modification as well as lipoate biosynthesis (143). Indeed, bioassays demonstrated that the low lipoyl protein content of the lipB null mutants was further depressed to undetectable levels in the lipB lplA double mutants (143). This finding suggested that the attenuated, but still detectable, accumulation of protein-bound lipoate by lipB null mutants was entirely due to the action of the lplA gene product. Moreover, overexpression of LplA allowed normal growth of lipB null mutant strains in the absence of lipoate, thus clearly demonstrating the redundant roles of these two genes (143). Genetic and biochemical evidence demonstrated that lipB encoded the octanoyl-ACP:protein N-octanoyltransferase (which is also an lipoyl-ACP:protein N-lipoyltransferase) (Fig. 9). An enzyme activity was detected in E. coli cell extracts that catalyzed the transfer of octanoic acid (lipoic acid) from octanoyl-ACP (lipoyl-ACP) to E2 apo-domains (Fig. 9). This activity was also found in extracts of E. coli lplA null mutants and, unlike LplA, this activity was not dependent on ATP. However, transferase activity was absent in E. coli lipB mutants (114). A temperature-sensitive lipB strain was obtained and found to encode a transferase of decreased thermal stability (113), indicating that lipB encoded the transferase rather than playing a regulatory role. Finally, His-tagged LipB was purified and the purified protein had high levels of octanoyltransferase and lipoyltransferase activities (113). The untagged protein has recently been purified by conventional means (145).
Based on these observations, a two-pathway E. coli lipoylation system was proposed (114, 143, 234) (Fig. 10). When presented with free lipoic acid in the medium, E. coli incorporates extracellular lipoate (98, 171) via the LplA-dependent scavenging pathway that utilizes ATP to activate lipoic acid in the form of lipoyl-AMP. When lipoate is absent from the medium, lipoyl groups must be made by de novo synthesis. An octanoyl group is first transferred from octanoyl-ACP to the apo-proteins by LipB. LipA then acts on the protein-bound octanoyl groups to catalyze the sulfur insertion step (Fig. 10). This model explains why lplA null mutants showed no growth defects unless the strain also carried a lipA or lipB mutation as well as the unidirectional redundancy between LipB and LplA functions. Indeed, LplA utilizes octanoyl-ACP as substrate with low but detectable efficiency (see below), accounting for the leakiness of lipB strains. It should be noted that strains having null mutations in both lplA and lipB contain no detectable lipoylated proteins, indicating that LplA and LipB are the only E. coli enzymes capable of modifying lipoyl domains (143).
Three assays have been used to detect attachment of lipoic acid to apo forms of PDH and 2-OGDH in vitro. The first assay measured the conversion of radioactive lipoate (or octanoate) to a protein-bound form defined as being insoluble in organic solvents (142). This is a sensitive and quantitative assay but applicable only to LplA since both the LipB substrate and product are protein bound. The second assay relies on the use of inactive unmodified apo-PDH or 2-OGDH complex purified from a lipB lplA strain completely deficient in modification of the E2 proteins. Lipoylation of the purified apo-PDH or 2-OGDH complex was detected by assay of the products of ligation reactions for either PDH or 2-OGDH activity (111). The third assay is a gel mobility shift assay (114). It follows the acylation-dependent shift in the electrophoretic mobility of a purified 87-residue apo-lipoyl domain from the E. coli PDH complex (7). This assay is much less sensitive than the other two but has the advantage that it can be used with any acyl donor because the mobility shift is due to loss of the positive charge of the lysine residue. With this assay it was found that purified LipB could convert the apo form of lipoyl domain to the holo domain with either octanoyl-ACP or lipoyl-ACP as the substrate. When LipB was tested for the ability to use ATP plus lipoic acid or octanoic acid, no modification was detected. The genetic studies discussed above suggested that LplA could catalyze the transferase reaction at a low rate (143). To test whether LplA possessed octanoyltransferase (lipoyltransferase) activity, the ability of purified LplA to use octanoyl-ACP or lipoyl-ACP as a substrate was tested in vitro. Gel mobility shift assays showed that LplA purified from a lipB null mutant strain did indeed use lipoyl-ACP and octanoyl-ACP as substrates, although it was extremely inefficient, a result consistent with the need to overproduce LplA in order to suppress a lipB mutant. A nonhydrolyzable lipoyl-AMP analogue was found to competitively inhibit both the ACP-dependent and the ATP-dependent reactions of LplA, indicating that the same active site catalyzed two chemically diverse reactions (113). How the LplA active site copes with such different chemistries remains a mystery, especially given that the LipB reaction proceeds via an acyl enzyme intermediate (see below). However, another enzyme reaction having an acyl enzyme intermediate proceeds normally upon elimination of the active-site residue that normally carries the intermediates (206). Hence, the geometry of the LplA active site might be sufficient to explain the low rates of octanoyl transfer from octanoyl-ACP.
Another lipB phenotype is that multiple copies of the gene confer resistance to selenolipoic acid. An analogue-resistant mutant that did not map at the lplA locus (172) was shown to be a chromosomal amplification of the lipA lipB region of the chromosome (112). The increased lipB dosage resulted in greater LipB activity that resulted in increased levels of lipoylation by endogenously synthesized lipoic acid that competed with the utilization of exogenous selenolipoic acid via the LplA-dependent pathway. A very modest (two- to threefold) increase in lipB dosage was sufficient to give resistance, which was explained by the known highly nonlinear relationship between the degree of protein lipoylation and the activity of the 2-oxoacid dehydrogenase complexes plus the fact that E. coli does not require full activity of the 2-oxoacid dehydrogenases for growth on minimal medium containing glucose (112). Thus, synthesis of sufficient lipoic acid to modify a few percent of the 2-oxo acid dehydrogenase complexes allowed growth in the presence of selenolipoic acid.
As mentioned above, the LipB reaction was recently shown to proceed via an acyl enzyme intermediate (233). The octanoyl group is transferred from the ACP thiol to the thiol of Cys-169. This protein-bound thioester is then attacked by the ε-amino group of the lipoyl domain lysine to give the modified domain.
Octanoyl-ACP + LipB ↔ [Octanoyl-LipB] + ACP-SH
[Octanoyl-LipB] + Apo-lipoyl domain → Octanoyl-E2 domain
The reactivity of the cysteine residue seems responsible for the only LipB crystal structure currently available, that of the Mycobacterium tuberculosis enzyme (134). The M. tuberculosis LipB, the expression of which was shown to complement growth of E. coli lipB mutant strains, was crystallized in a covalent complex with decanoic acid. Surprisingly, although the acyl chain was bound to the sulfur atom of a cysteine residue corresponding to Cys-169 of E. coli LipB, the bond was a thioether linkage to C-3 of decanoate rather than a thioester link to the carboxyl group (134). This unexpected finding seems likely to be the result of a Michaelis addition of the cysteine thiol to the unsaturated bond of trans-2 decenoyl-ACP or cis-3-decenoyl-ACP, a key intermediate in E. coli unsaturated fatty acid biosynthesis. Consistent with this interpretation, no such adduct was seen upon expression of the protein in Mycobacterium smegmatis, which forms unsaturated fatty acids by a pathway that does not involve decenoyl intermediates (134). Based on this crystal structure and mutagenesis studies, LipB is thought to function as a novel cysteine/lysine dyad acyltranferase, in which the dyad residues function as acid/base catalysts (134). Structural comparison with LplA rationalizes the finding that LplA is able to catalyze the LipB reaction since the two enzymes have remarkably similar active-site geometries and quite similar overall structures (134).
Although the functions of lipoic acid in the multienzyme complexes have been well studied over the past forty years, an understanding of the lipoic acid biosynthesis pathway has been achieved only recently. Such studies have focused on E. coli. Early studies had established that octanoic (properly n-octanoic) acid (Fig. 1) is the precursor of the lipoic acid carbon chain (157). Analysis of the conversion of specifically labeled forms of octanoic acid to lipoic acid by E. coli cultures showed that sulfur atoms are introduced with loss of only two hydrogen atoms from the chain, one from C-6 and the second from C-8 (155, 219). Additional metabolic feeding studies demonstrated that E. coli lipoic acid biosynthesis does not involve either desaturation or hydroxylation of octanoic acid but does result in inversion of stereochemistry at C-6 (155, 156). Sulfur is introduced at C-8 with racemization in agreement with the formation of an intermediate carbon radical at C-8 (156, 218, 219, 220). 8-Thiooctanoic acid and 6-thiooctanoic acid were readily converted to lipoic acid, although 6-thiooctanoic acid was converted only 10 to 20% as efficiently as the other positional isomer (218). Genetic studies identified a single E. coli gene responsible for the sulfur-insertion steps of lipoic acid biosynthesis, first called lip (97) and now called lipA, which encodes a protein called lipoic acid synthase. E. coli strains having null mutations in lipA do not synthesize lipoic acid, and the phenotypes of these mutants suggested that LipA was responsible for the formation of both C–S bonds (94, 95, 171, 211) which encodes a protein called lipoic acid synthase. E. coli strains having null mutations in lipA do not synthesize lipoic acid, and the phenotypes of these mutants suggested that LipA was responsible for the formation of both C–S bonds (97) and encodes a protein now called lipoic acid synthase.
There are strong parallels between LipA and biotin synthase (BioB), the enzyme discussed above that catalyzes the final step in the biotin biosynthesis (Fig. 4). LipA-like BioB makes two C–S bonds and also removes two unactivated hydrogen atoms. The similarity in chemistry between the biosynthesis of the dithiolane ring of lipoate and the thiophane ring of biotin strongly suggests functional parallels in the mechanisms of the enzymes that produce these compounds. Indeed, the amino acid sequences of the E. coli LipA and BioB proteins show marked similarities; 40% sequence similarity and 17% sequence identity (171).
As discussed above, BioB has both a [4Fe-4S] cluster and a [2Fe-2S] cluster. The canonical iron-sulfur cluster binding motif CXXXCXXC is also found in the LipA sequence leading to early predictions that it is an iron-sulfur protein (94, 171). The LipA protein has been overexpressed in E. coli and purified from both soluble lysates and insoluble aggregates that were subsequently refolded and reconstituted with ferrous iron and sulfide (33, 149, 171). The purified dimeric protein (33) has a dark reddish-brown color and displays a band at 420 nm in its light absorption spectrum, characteristic of a sulfide-to-iron charge transfer. Resonance Raman, electronic absorbance, and Mössbauer spectroscopic results were consistent with the presence of an iron-sulfur cluster in LipA. It was suggested that LipA contains [2Fe-2S] clusters that during reduction are converted into [4Fe-4S] clusters (33, 149). The Fe-S cluster of LipA was first suggested to be a [4Fe-4S] cluster bridging the two subunits (33). However, in a different report it was suggested that the limited amount of Fe and S atoms and the presence of [2Fe-2S] clusters in the previous preparation of LipA were a direct consequence of aerobic isolation. It was reported that under strictly anaerobic conditions LipA could bind one [4Fe-4S] cluster per subunit (150). Recently, it was reported that LipA contains two distinct [4Fe-4S] clusters per polypeptide (46). Thus, the types of disagreements seen in the BioB literature are also apparent for LipA, showing the obvious difficulties of working with this class of proteins.
Direct involvement of LipA in the sulfur insertion reaction of lipoic acid biosynthesis was difficult to establish because of the lack of an in vitro assay. Much of this difficulty was due to the assumption that free octanoic acid was the sulfur acceptor. The first indication that this was not the case was the demonstration by Jordan and Cronan (114) of the LipB transferase activity. Miller and coworkers (141) were the first to report synthesis of lipoic acid in vitro. This was based on the discovery of LplA and LipB that led to development of a defined in vitro lipoic acid synthesis system and an assay that was much more sensitive and quantitative than prior assays (141). Lipoic acid synthesis was assayed indirectly using (i) the apo form of pyruvate dehydrogenase complex (apo-PDH) as a lipoyl-accepting protein, (ii) purified LipA, and either (iii) purified LplA, ATP, octanoic acid as a substrate (for lipoic acid synthesis) or (iv) LipB and octanoyl-ACP as a substrate. Activation of apo-PDH upon lipoylation was monitored spectrophotometrically via reduction of an NAD+ analogue. Within a finite range, the rate of reduced pyridine dinucleotide formation directly depended on the amount of lipoylated PDH. This assay showed that LipA is responsible for both of the sulfur insertions and that octanoyl-ACP (or a derivative of octanoyl-ACP), but not octanoic acid, was a LipA substrate. Moreover, this work showed that, as suspected, the LipA reaction requires iron-sulfur clusters and SAM to perform the radical chemistry. The principal disadvantage of this assay was its indirect nature and detection of lipoylation of apo-PDH rather than of the primary lipoyl protein species per se. All attempts to isolate a free lipoyl-ACP product in the assay were unsuccessful. Thus, the exact identity of the immediate product of the LipA reaction could not be determined by this assay. Recent studies demonstrate that LipA acts on octanoylated derivatives of lipoyl-accepting proteins (25, 234).
The first evidence that octanoyl domain rather than octanoyl-ACP was the substrate for sulfur insertion was the finding that lipB mutants grew well when supplemented with octanoic acid in place of lipoic acid (234). Octanoate supplementation of lipB strains required function of both the lipA and lplA genes; both lipB lipA and lipB lplA doubly mutant strains failed to grow on octanoate. Moreover, growth was specific to octanoate, and fatty acids of 6, 7, 9, and 10 carbons were inactive (234). These observations argued for the existence of an LplA-dependent pathway that bypassed LipB function in the presence of octanoate. In the postulated bypass pathway (Fig. 10), LplA would attach octanoate derived from the growth medium to the unmodified E2 domains of the PDH and 2-OGDH E2 subunits. LipA would then insert two sulfur atoms into the covalently bound octanoyl moiety and thereby convert the octanoyl-E2 domains to lipoyl-E2 domains in situ. That is, lipoic acid would be assembled on its cognate proteins. The resulting active enzymes would account for the observed growth of lipB strains on octanoate (Fig. 10). This pathway was tested in vivo (234). First, an 87-residue E2 domain derived from E. coli PDH was expressed in a host strain that carried null mutations in lipA (to prevent lipoic acid synthesis), lipB (to block octanoate transfer from fatty acid synthesis), and fadE (to block β-oxidative degradation of octanoate). The use of the domain allowed detection of modification by the electrophoretic mobility shift assay and by mass spectroscopy. When this strain was cultured in a medium supplemented with octanoic acid about half of the domain became modified. In addition, the LipB-dependent modification pathway was assayed in a lipA lplA null mutant strain grown in the absence of exogenous octanoate. In agreement with prior work using a lipA strain (7), octanoyl-E2 domain accumulation was detected. Therefore, the E2 domain could be octanoylated in vivo either by LplA using exogenously added octanoate or by LipB using de novo synthesized octanoate. In order to assay conversion of octanoyl-E2 domain to lipoyl-E2 domain the lipA lipB fadE strain was supplemented with deuterated octanoic acid to allow accumulation of octanoyl d15-E2 domain that was readily distinguished by mass spectroscopy from domain modified with endogenously synthesized octanoate. Following accumulation of the d15-labeled octanoyl-E2 domain LipA function was restored by transduction with cells with phage λ particles containing a lipA cosmid that had been packaged in vivo. By using this approach, two types of labeling experiments were done. In the first protocol E2 domains were labeled in vivo by growth in the presence of octanoic d15 acid. Following removal of the labeled octanoate the cells were then resuspended in growth medium and transduced with the packaged lipA-encoding cosmid. Following incubation to allow lipoate synthesis, samples were taken and the E2 domain species were isolated, purified, and analyzed by electrospray mass spectroscopy (Fig. 11). In the cultures to which LipA activity was restored a readily detectable conversion of the E2 domain modified with octanoate d15 to a species of 60 additional mass units was seen. This was exactly the increase in mass (gain of two sulfur atoms of mass 32 and loss of two deuterium atoms of mass 2) expected for conversion of the d15-labeled octanoyl-E2 domain to the d13-labeled lipoyl domain. In the second protocol (a variation of the first protocol) the octanoic d15 acid was removed by washing the cells and replaced with normal (nondeuterated) octanoate. This experiment gave essentially the same result; the d15-labeled octanoyl-E2 domain was converted to d13-labeled lipoyl-E2 domain (Fig. 11). A modification of these experiments also showed that octanoyl-PDH accumulated in vivo in a lipA strain was converted to its active form upon restoration of LipA activity (234). The conversion of octanoyl domain to lipoyl domain was also observed in vitro (234), although the extent of conversion was much less than stoichiometric with LipA. These results were recently confirmed by using octanoyl-H protein as the substrate with an eightfold increase in the yield of lipoic acid formed/LipA monomer (45).
As mentioned above, lipoic acid synthase is a member of the radical-SAM enzyme superfamily that utilizes a reduced iron-sulfur cluster and SAM to generate 5'-deoxyadenosyl (DOA) radicals for further radical-based chemistry (40, 74, 108, 146, 189). In the lipoic acid synthase reaction (Fig. 12), it is generally believed that the role of the DOA radical is to remove one hydrogen atom from each of the C-6 and C-8 positions of octanoic acid, thereby allowing for subsequent sulfur insertion (45, 141). Consistent with this prediction two molecules of SAM are required to synthesize one mole of lipoyl cofactor (45). This stoichiometry is similar to that obtained in the two studies in the BioB reaction (91, 185) and suggests that the abortive cleavage of SAM observed in these systems might result from some innate reactivity associated with this subclass of radical SAM enzymes (45).
As in the case of biotin, the source of the sulfur atoms of lipoic acid is thought to be a [Fe-S] cluster distinct from the SAM radical [4Fe-4S] cluster. In the first successful in vitro lipoic acid synthesis assays, lipoic acid was formed in the absence of exogenous sulfur-containing compounds in the in vitro assay (45, 141). This suggested that, like biotin synthase, the protein itself has some mobilizable sulfur atoms, either from an Fe-S cluster, a persulfide, or some other species. The currently favored sulfur source is an iron-sulfur cluster (137, 141). Recent work from the Booker group reported that lipoic acid synthase contains two [4Fe-4S] clusters (46). One cluster is coordinated by the radical SAM CXXXCXXC motif that functions in DOA radical generation. The second cluster is coordinated by the CXXXXCXXXXXC motif, which, thus far, is unique to lipoic acid synthases and suggested to be the source of the sulfur atoms. This has been addressed (as in BioB; 81) by preparation of LipA from E. coli cells grown on an isotopically labeled sulfur source (34S) (44). As expected, the lipoic acid formed using this enzyme in vitro was isotopically labeled with 34S. Moreover, when the reactions were performed with equimolar amounts of 32S-labeled LipA and 34S-labeled LipA, the lipoic acid molecules formed contained either two 32S atoms or two 3S atoms (44). Thus, both sulfur atoms emanate from the same polypeptide, thereby eliminating the possibility that the monothiolated species are released and the second sulfur atom is inserted following rebinding to LipA. If release and rebinding occurred, half of the lipoic acid formed would have one atom each of 32S and 34S. Such mechanistic experiments will be facilitated if an octanoylated peptide substrate can be substituted for the octanoylated lipoyl domain as in the case for an octanoyl tripeptide in the Sulfolobus solfataricus LipA assay (30). This system has recently been used to show that S. solfataricus LipA catalyzed lipoate biosynthesis in a stepwise manner, with sulfur first being inserted at C-6 of the octanoyl chain (68). However, this intermediate remained tightly bound to LipA, consistent with the finding that both sulfur atoms are derived from the same LipA polypeptide. However, as first noted by Parry (155), there is a large isotope effect at C-8 that complicates interpretation of such labeling studies. A current model of the LipA reaction is given in Fig. 12.
It should be noted that the finding that lipoic acid synthesis proceeds through an octanoyl-domain intermediate explains a previously puzzling observation first made by Ali and Guest (6, 7) and subsequently by others (100). These workers found that, upon overproduction of a lipoyl domain in E. coli, three species of domain were obtained, the expected apo and lipoylated domains plus a third species subsequently shown to be octanoylated domain. Based on conventional biochemistry in which LipA would produce free lipoic acid, the octanoylated domain was thought to be an anomalous product resulting from the lack of sufficient free lipoic acid to modify the overexpressed domain plus a lack of specificity of the attachment enzyme (6, 7). From the present pathway it now seems clear that LipA was limiting (as was proposed), but the octanoylated domain was an accumulated intermediate rather than the aberrant by-product of overproduction.
As in the case of BioB, LipA has not been shown to be catalytic in vitro; the best preparations to date form only about 0.4 molecules of lipoic acid per LipA molecule, and thus the sacrificial protein scenario of BioB might pertain to this protein. The question of whether or not LipA is catalytic in vivo remains to be tested. If results analogous to those of BioB are obtained, the question is how the novel LipA CXXXXCXXXXXC motif becomes liganded into a [4Fe-4S] cluster. In this regard, coexpression of LipA with the Isc proteins results in increased LipA activity (123). However, it remains to be seen which clusters are built by this manipulation. The crystal structure of LipA with SAM and octanoylated domain would be of great benefit. This would be facilitated if an octanoylated peptide substrate can be substituted for the octanoylated lipoyl domain, as is the case for S. solfataricus LipA (30). It has been predicted that LipA is a (α6β6) barrel protein (a three-quarters barrel) rather than a full (α6β6) TIM barrel like BioB (146). Moreover, LipA is reported to contain two distinct [4Fe-4S] clusters, whereas BioB is believed to contain one [4Fe-4S] cluster and one [2Fe-2S] cluster. Therefore, although many of the questions that have bedeviled the BioB literature such as the sulfur source are germane to LipA, it seems clear that there are significant differences in how the two enzymes accomplish their reactions. A LipA crystal structure would be most useful in understanding these differences.
Another major question concerns the regulation of lipoic acid synthesis. Although the lipB and lipA genes lie close to one another on the E. coli chromosome and are transcribed in the same direction, the genes are separated by 1.4 kbp and this spacer region contains ybeF, an open reading frame that encodes a possible LysR-type transcription factor. Strains carrying transposon insertions into and deletions of ybeF have no phenotype indicating that lipB and lipA are not in an operon (12, 211). Is expression of these genes regulated? Lipoic acid is clearly synthesized during aerobic growth, and anaerobic function of the glycine cleavage enzyme indicates that it is also made under fermentative conditions (211). Moreover, a recent report that E. coli contains high levels of 2-oxoglutarate dehydrogenase when grown anaerobically with an electron acceptor such as nitrate (163) indicates that lipoic acid synthesis must also proceed under these growth conditions. The transcription of lipA in S. enterica has been assayed by transcriptional fusions to β-galactosidase and was found to be unaffected by catabolite repression conditions or by addition of lipoic acid (188). Therefore, the lipoic acid synthesis pathway may be constitutively expressed. Given the unusually sophisticated bio operon transcriptional regulatory system, it might seem unlikely that lipoic acid synthesis is unregulated. Indeed, biotin and lipoic acid are synthesized at similar levels in E. coli. However, biotin synthesis requires six enzymes and several of the reactions of the pathway require input of metabolically expensive molecules, which might justify regulation of biotin synthesis enzyme production. In contrast, octanoate, the precursor of the lipoic acid carbon chain, is derived from fatty acid biosynthesis in an already activated form, octanoyl-ACP, and lipoate synthesis consumes only a tiny fraction of the total cellular fatty acid synthetic capacity. Since LipB uses a preformed activated intermediate, the only further energetic input (in the form of SAM) occurs in the LipA sulfur insertion reaction. Therefore, relative to biotin synthesis, lipoic acid synthesis has a low metabolic price. Another consideration is that the lipoic acid synthesis pathway is limited by the amount of apo-lipoyl domain available and thus, unlike biotin synthesis, cannot "run wild" to overproduce and excrete the cofactor as is the case when the bio operon is deregulated (36). Thus, it can be reasonably argued that regulation of the lipoic acid synthetic pathway might well be more expensive than the alternative of simply allowing constitutive expression of the genes. A perplexing observation is the report that LipB acts as a negative regulator of deoxyadenosine methyltransferase (dam) gene expression in E. coli (209). These workers speculate that LipB may inactivate a repressor protein by lipoylation. However, all of the proteins that become labeled with exogenous radioactively labeled lipoate or octanoate in vivo are known subunits of the enzymes discussed above (142, 211). Hence, the putative lipoylated repressor would have to be modified by LipB, but not by LplA. Further studies of this interesting phenomenon are needed.
It is gratifying that the recent crystal structures have resulted in LplA, LipB, and BirA being recognized as a new protein family (PFAM 03099.13) as predicted by Reche (169), although these proteins share only a single conserved residue. Indeed the Cα carbons of E. coli LplA and BirA (minus the DNA binding domain) structures can be aligned with a root-mean-square deviation of 2.8 Å over much of their lengths (227), whereas E. coli LplA and M. tuberculosis LipB can be aligned over the length of LipB (the smaller protein) with a Cα value of 2.5 Å (134). Moreover, the cofactor ligands in these crystal structures are in register indicating similar geometries of binding. These findings together with the even more similar structures of the domains that are the substrates of these enzymes raises the question of how the cell avoids the catastrophic effects on metabolism that would result if biotinylated proteins became lipoylated and vice versa. This question of accurate modification has been addressed by Reche and Perham (167, 168), who showed that the wild-type biotinoyl domain can be lipoylated by LplA in vitro, but the reactions proceed only with a molar excess of enzyme and very slowly. They also showed that mutations of the residues adjacent to the lysine residue of the biotinoyl domain (the protruding β-turn) to those found in the protruding β-turn of the lipoyl domain allow modification by lipoic acid attachment. Indeed, conversion of a single residue of the biotinoyl domain of E. coli allowed some lipoylation by LplA. Moreover, some of the hybrid domains remained substrates for biotinylation. However, these workers also found that they could leave the protruding β-turn region intact and obtain lipoylation by removal of the biotinoyl domain thumb structure that is not found in lipoyl domains (167, 168) (Fig. 8). Thus, the thumb structure seems to be the major "gate-keeper" that prevents lipoylation of AccB. This seems an appropriate choice since the thumb structure is required for function of AccB in fatty acid synthesis, but not for biotinylation of the protein (51). The determinants that prevent biotinylation of lipoyl domains have yet to be explored.
In the two print edition predecessors of EcoSal, the biotin and lipoic acid synthetic pathways were placed into the same chapter. It seems probable that this was because both are sulfur-containing and covalently attached enzyme cofactors, plus there was insufficient information on lipoic acid synthesis to justify a separate chapter. However, in the 11 years that have elapsed since the second edition, the lumping of these cofactors together seems prescient. We now know that there are many commonalities. Both cofactors are made as offshoots of the fatty acid synthetic pathway, sulfur insertion is done by SAM radical enzymes, the sulfur is almost certainly derived from an enzyme iron-sulfur center, the cofactors are attached to very similar domains, and the enzymes that attach these cofactors constitute a protein family despite the low sequence conservation and differing mechanisms of the enzymes.
The preparation of the manuscript and the experimental work from our laboratory were supported by Grant AI15650 from the National Institute of Allergy and Infectious Diseases.
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Module3.6.3.5
