A Trail of Research from Lipoic Acid to α-Keto Acid Dehydrogenase Complexes (2001)

pyruvate dehydrogenase α-ketoglutarate dehydrogenase binding protein truncated E2 In this article I shall retrace a trail of research that began with the isolation and characterization of a microbial growth factor and led to elucidation of the structure, function, and regulation of α-keto acid dehydrogenase complexes. The high points of this trail are presented below. This trail of discovery started in the spring of 1949, about 6 months after I joined the faculty of the Department of Chemistry at the University of Texas. At that time I started working on the isolation of a factor that replaced acetate in the growth medium for certain lactic acid bacteria. Research on the “acetate-replacing factor” was initiated by Esmond Snell and associates at the University of Wisconsin and then at the University of Texas. I inherited this project in the spring of 1949. We established that this factor is widely distributed in animal, plant, and microbial cells and that liver is a rich source. The factor is tightly bound to liver protein and is released by proteolysis or by acid hydrolysis. At that time pharmaceutical companies were processing large amounts of pork and beef liver to obtain extracts suitable for treatment of pernicious anemia. The active principle was shown later to be vitamin B12. Fresh liver was extracted with warm water, and the residual liver proteins and fatty material were dried and sold as an animal feed supplement. Arrangements were made with Eli Lilly and Co. to obtain liver residue, and we developed procedures for extracting and purifying the acetate-replacing factor. We progressed to the point of being able to process about 6 pounds of liver residue at a time. A 16,000–50,000-fold purification was achieved. In the late 1940s and early 1950s several other groups were trying to isolate factors that were similar to, if not identical with, the acetate-replacing factor. These factors included the “pyruvate oxidation factor” of I. C. Gunsalus and associates that was necessary for oxidation of pyruvate to acetate and carbon dioxide byStreptococcus faecalis cells grown in a synthetic medium. Gunsalus was also collaborating with Eli Lilly and Co. In the fall of 1950, the Lilly Research Laboratories merged the two separate collaborations to facilitate isolation of the acetate-replacing/pyruvate oxidation factor. The Lilly group adapted and scaled up isolation procedures developed by us. Instead of processing 6-pound batches of liver residue at a time, they were able (using commercial equipment) to process 250-pound batches. Concentrates of the factor that were 0.1–1% pure were sent to my laboratory for further processing. I obtained the first pale yellow crystals of the factor, about 3 mg, on or about March 15, 1951, a truly memorable occasion. It was partially characterized and given the trivial name α-lipoic acid (1Reed L.J. DeBusk B.G. Gunsalus I.C. Hornberger Jr., C.S. Crystalline α-lipoic acid: a catalytic agent associated with pyruvate dehydrogenase..Science. 1951; 114: 93-94Crossref PubMed Scopus (224) Google Scholar). The isolation involved a 300,000-fold purification. A total of ∼30 mg of crystalline lipoic acid was eventually isolated. We estimated that ∼10 tons of liver residue were processed to obtain this small amount of the pure substance. And to think that I was processing about 6 pounds of liver residue at a time, convinced that I would eventually isolate the pure material. We established that lipoic acid is a cyclic disulfide, either 6,8-, 5,8-, or 4,8-dithiooctanoic acid. That the correct structure is 6,8-dithiooctanoic acid (1,2-dithiolane-3-valeric acid) was established by synthesis of dl-lipoic acid, first achieved by E. L. R. Stokstad and associates at Lederle Laboratories. I was intrigued by this simple yet unique substance and wanted to know more about its biological function, i.e. with what and how does it function in living cells. Little did I know then that this trail would take me through five decades of research that has turned out to be a fascinating and rewarding chapter of modern biochemistry, elucidation of the mechanism of oxidative decarboxylation of α-keto acids. Prior to the isolation of lipoic acid and its characterization as a cyclic disulfide, contributions from several laboratories had established the cofactor requirements for the oxidative decarboxylation of α-keto acids represented by the equation shown below. RCOCO2−+CoASH+NAD+→RCOSCoA+CO2+NADHIn addition to CoA and NAD+, thiamin diphosphate, a divalent metal ion, and protein-bound lipoic acid are required. A requirement for FAD was demonstrated later. The presence of a disulfide linkage in lipoic acid led Gunsalus to propose that lipoic acid underwent a cycle of reactions in α-keto acid oxidation comprising reductive acylation, acyl transfer, and electron transfer. Lipoic acid was visualized as functioning after thiamin diphosphate and before CoA and NAD+. Gunsalus, Lowell Hager, and associates obtained evidence for this proposal using lipoic acid and derivatives thereof in substrate amounts. However, the physiological reactions presumably involve catalytic amounts of protein-bound lipoic acid. Elucidation of the nature of the functional form of lipoic acid was essential for verification of the postulated reactions and for further clarification of mechanism. We decided to focus our attention initially on S. faecalis (strain 10C1), which had been grown on a lipoic acid-deficient medium. As shown by Gunsalus and associates these cells did not oxidize pyruvate or α-ketobutyrate unless lipoic acid was added to the preparations. However, attempts to activate cell-free extracts of the lipoic acid-deficient cells with lipoic acid or natural extracts containing complex forms of lipoic acid were unsuccessful. We discovered that cell-free extracts of the deficient cells could be activated by incubation with lipoic acid prior to addition of substrate and supplements. Approximately 30 min of preincubation were required for maximal activation. Activity was reduced only slightly by dialysis, suggesting that lipoic acid was converted to the enzymatically active, “bound” form during the incubation. That such is the case was shown by experiments with lipoic acid-35S2. By fractionation of the lipoic acid-deficient extracts, we determined the components required and the nature of the reactions involved in activation of the extract by lipoic acid. One fraction contained the apopyruvate oxidation system, and a second fraction contained a lipoic acid-activating system. A requirement for ATP was established, suggesting that lipoic acid was activated through its carboxyl group before incorporation into the apopyruvate oxidation system. Based on the results of Paul Berg and others at that time, demonstrating that acyl adenylates are produced by enzyme-catalyzed interaction of organic acids and ATP, we demonstrated that lipoic acid and ATP could be replaced by synthetic lipoyl adenylate. The lipoic acid-activating system, i.e. lipoate-protein ligase, was also detected inEscherichia coli and partially purified. During the course of these studies with cell-free extracts of lipoic acid-deficient S. faecalis, several amides of lipoic acid were synthesized for testing as possible antagonists of the lipoic acid-activating system. These amides did not inhibit activation of the cell-free extract by lipoic acid. On the contrary, these amides could replace lipoic acid, provided ATP was present. These observations suggested that hydrolysis of the amides occurred during incubation with the cell-free extract or the partially purified enzyme preparations. The hydrolytic enzyme, designated lipoyl X-hydrolase and later shown to be a lipoamidase, was purified about 100-fold. The availability of lipoamidase facilitated our preliminary studies in the late 1950s on the biosynthesis of lipoic acid in E. coli. Possible radioactive precursors were included in the growth medium. The PDH1 and KGDH complexes were purified and treated with lipoamidase. The released lipoic acid was extracted into benzene and its radioactivity was determined. We found that octanoate-1-14C was incorporated into lipoic acid as a unit, C-1 of lipoic acid corresponding to C-1 of octanoic acid. Recently, Michael Marletta, John Cronan, and associates have shown that lipoyl synthase, which contains an iron-sulfur cluster, catalyzes the insertion of sulfur at C-6 and C-8 of octanoyl-acyl carrier protein to produce lipoyl-acyl carrier protein. Lipoamidase and lipoate-protein ligase proved to be invaluable in providing direct, unequivocal evidence of the involvement of protein-bound lipoic acid in the CoA- and NAD+-linked oxidative decarboxylation of pyruvate and α-ketoglutarate and in providing clarification of the mechanism of model reactions catalyzed by the pyruvate and α-ketoglutarate dehydrogenase complexes and components thereof. This evidence comprised a demonstration of inactivation and reactivation of the enzyme or enzyme complex accompanying, respectively, release and reincorporation of the lipoyl moiety. When E. coli (Crookes strain) was grown aerobically in the presence of lipoic acid-35S2, the radioactive substance was incorporated into the pyruvate and α-ketoglutarate dehydrogenation systems, due to the presence of lipoate-protein ligase. The availability of the highly purified complexes permitted rapid progress in the late 1950s in identification of the moiety to which lipoic acid is bound. The protein-bound radioactive lipoyl moiety was oxidized with performic acid, and the protein was partially hydrolyzed with 12 n hydrochloric acid (3 h at 105 °C). From the hydrolysates Hayao Nawa (2Nawa H. Brady W.T. Koike M. Reed L.J. Studies on the nature of protein-bound lipoic acid..J. Am. Chem. Soc. 1960; 82: 896-903Crossref Scopus (52) Google Scholar) isolated in good yield a ninhydrin-positive, radioactive conjugate, which was identified as ε-N-(6,8-disulfooctanoyl)-l-lysine by degradation and synthesis. The lipoyl moiety in the two complexes therefore is bound in amide linkage to the ε-amino group of a lysyl residue (Fig. 1). Prior to 1950 pyruvate and α-ketoglutarate oxidation had been studied mainly with particulate preparations that were unsuitable for detailed analysis. Solubilization of bacterial and animal α-keto acid oxidation systems in the early 1950s in the laboratories of Severo Ochoa and David Green was a significant advance. Seymour Korkes, Gunsalus, and Ochoa succeeded in separating the pyruvate oxidation system of anaerobically grown E. coli (strain ATCC 4157) into two components, designated Fraction A and Fraction B. Subsequently, Hager and Gunsalus found that extracts of aerobically grown E. coli (Crookes strain) contained 30–50 times the pyruvate oxidation activity of the anaerobically grown 4157 cells. Using lipoic acid and dihydrolipoic acid in substrate amounts they showed that Fraction A contained a lipoyl transacetylase and that Fraction B contained a lipoyl dehydrogenase. Richard Schweet and associates isolated a CoA- and NAD+-linked pyruvate oxidation system from pigeon breast muscle in a highly purified state, with an apparent molecular weight of about 4 million. D. R. Sanadi and associates isolated a CoA- and NAD+-linked α-ketoglutarate oxidation system from pig heart with an apparent molecular weight of 2 million. In my laboratory we developed mild procedures for purification of the pyruvate and α-ketoglutarate oxidation systems from E. coli (Crookes strain). By the late 1950s Masahiko Koike (3Koike M. Reed L.J. Carroll W.R. α-Keto acid dehydrogenation complexes. I. Purification and properties of pyruvate and α-ketoglutarate dehydrogenation complexes of Escherichia coli..J. Biol. Chem. 1960; 235: 1924-1930Abstract Full Text PDF PubMed Google Scholar) succeeded in isolating these enzyme systems as highly purified functional units with molecular weights in the millions. It was very exciting to see in the analytical ultracentrifuge of my friend and collaborator at NIH, William Carroll, a major symmetrical peak for each of the two highly purified preparations and that the boundary of the yellow color of the flavoprotein was associated with the main peak. The molecular weights of these multienzyme units were determined to be 4.8 and 2.4 million, respectively. By careful and persistent work over a period of several years, we dissected the pyruvate and α-ketoglutarate dehydrogenase complexes into their component enzymes, characterized them, and reassembled the large functional units from the isolated enzymes (4Koike M. Reed L.J. Carroll W.R. α-Keto acid dehydrogenation complexes. IV. Resolution and reconstitution of the Escherichia coli pyruvate dehydrogenation complex..J. Biol. Chem. 1963; 238: 30-39Abstract Full Text PDF PubMed Google Scholar). We demonstrated that the individual enzymes are linked in the two complexes by non-covalent bonds and that by proper selection of experimental conditions the enzymes could be separated from one another without loss of enzymatic activity. We showed that each of these functional units is composed of multiple copies of three enzymes, a pyruvate and an α-ketoglutarate decarboxylase-dehydrogenase (E1), a dihydrolipoamide acetyltransferase and a succinyltransferase (E2), and a flavoprotein, dihydrolipoamide dehydrogenase (E3). These three enzymes, acting in sequence, catalyze the reactions shown in Fig.2. E1 catalyzes both the decarboxylation of the α-keto acid (Reaction 1) and the subsequent reductive acylation of the lipoyl moiety, which is covalently bound to E2 (Reaction 2).E2 catalyzes the acyl transfer to CoA (Reaction 3), and E3 catalyzes the reoxidation of the dihydrolipoyl moiety with NAD+ as the ultimate electron acceptor (Reactions 4 and 5). Binding experiments showed that the pyruvate dehydrogenase (E1) and the flavoprotein (E3) do not combine with each other, but each of these components does combine with the acetyltransferase (E2). The acetyltransferase serves a dual function, a catalytic function and a structural function,i.e. a scaffold for binding and localizingE1 and E3. In dilute acetic acid (0.83 m, pH 2.6) the acetyltransferase dissociated into inactive subunits with a molecular weight of about 70,000. Dilution of the acidic solution into suitable buffers resulted in restoration of enzymatic activity and the characteristic structure of the native acetyltransferase unit. The acetyltransferase appeared to be a self-assembling system. The two flavoproteins (from the PDH and KGDH complexes) were shown to be interchangeable with respect to both complex formation and function, and enzymatic, physical, and immunochemical data indicated that the two flavoproteins were very similar if not identical. It was evident that the lipoyl moiety undergoes a cycle of transformations, i.e. reductive acylation, acyl transfer, and electron transfer, involving three separate enzymes within a complex in which movement of the individual enzymes is restricted and from which intermediates do not dissociate. A possible molecular basis of these interactions emerged from our discovery that the lipoyl moiety is bound in amide linkage to the ε-amino group of a lysyl residue in the E2 component of the PDH and KGDH complexes. This linkage provides a flexible arm, about 14 Å in length (Fig. 1), conceivably permitting the lipoyl moiety to rotate among the active sites of E1, E2, andE3, i.e. a “swinging arm” active site coupling mechanism. Some 15 years later, spin label experiments by Richard Perham and Cees Veeger and their associates provided evidence that the lipoyllysyl residues are essentially free to rotate in the PDH complex. Our subsequent finding (see below) that the lipoyllysyl moiety is part of a “super arm,” i.e. a lipoyl domain, led to the proposal that movement of lipoyl domains as well as rotation of lipoyl moieties may provide the means to span the physical gaps between catalytic sites on E1,E2, and E3, as well as facilitate communication between lipoyl moieties. These were exciting times for us in the late 1950s and early 1960s. Our concept of the macromolecular organization of the PDH complex that emerged from these biochemical studies is that of an organized mosaic of enzymes in which each of the component enzymes is uniquely located to permit efficient coupling of the individual reactions catalyzed by these enzymes. This concept was confirmed and extended by electron microscopy studies conducted by my associate Robert Oliver. Electron micrographs of the E. coli PDH complex and its component enzymes negatively stained with phosphotungstate revealed that the complex had a polyhedral structure with a diameter of about 300 Å, that the acetyltransferase (E2) occupied the center of the polyhedron, and that the molecules of E1 andE3 were distributed on its surface. The shape of the acetyltransferase indicated that it had a cubelike structure. The shape of the succinyltransferase component of the E. coliKGDH complex was very similar. These results, together with biochemical data, demonstrated that both E2s consist of 24 apparently identical polypeptide chains arranged as eight trimers (morphological subunits) at the vertices of a cube (Fig.3A). This proposed structure was confirmed later by x-ray diffraction analyses carried out by collaborators David DeRosier and Marvin Hackert demonstrating that both acyltransferases possess 432 molecular symmetry. Our interpretative model of the macromolecular organization of the E. coli PDH complex in the mid-1960s depicted 12E1 dimers and 6 E3 dimers arranged, respectively, on the 12 edges and in the 6 faces of the cubelike E2. All dihydrolipoamide acyltransferases possess a unique multidomain structure. This architectural feature was revealed initially in my laboratory in the late 1970s by limited proteolysis studies of the E. coli dihydrolipoamide acetyltransferase containing [2-3H]lipoyl moieties. Dennis Bleile (5Bleile D.M. Munk P. Oliver R.M. Reed L.J. Subunit structure of dihydrolipoyl transacetylase component of pyruvate dehydrogenase complex from Escherichia coli..Proc. Natl. Acad. Sci. U. S. A. 1979; 76: 4385-4389Crossref PubMed Scopus (82) Google Scholar) found that limited tryptic digestion at pH 7.0 and 4 °C cleaved theE2 subunits (Mr∼64,500) into two large fragments, an outer lipoyl-bearing domain and an catalytic and binding The had a structure, and it the binding sites of the the binding sites for E1 andE3, and the catalytic site for transfer. The of catalytic and binding domains the of the the cubelike of this E2 with the electron The other tryptic designated the lipoyl domain, contained the covalently bound lipoyl moieties and had an extended structure. We suggested that the two domains are by a and that movement of lipoyl domains and not rotation of lipoyllysyl moieties may provide the means to span the physical gaps between catalytic sites on the complex. These early on the domain structure of dihydrolipoamide acyltransferases were confirmed and extended by studies involving molecular limited and in the laboratories of John and Richard the part of the acyltransferases contains or three highly similar lipoyl each of about acid The lipoyl domain is by another that is involved in binding These domains are linked to each other and to the part of the polypeptide by flexible that are rich in and acid These are to provide to the lipoyl active site coupling within the multienzyme complexes. In the late part of our research was isolation and characterization of the PDH and KGDH which are to within the were developed for of on a large from and heart the and of my friend and and mild procedures were developed to isolate the PDH and KGDH complexes from the In the course of attempts to these complexes in extracts of that the PDH but not the KGDH underwent a inactivation in the presence of A revealed that the and heart PDH complexes are by a cycle Reed L.J. α-Keto acid dehydrogenase complexes. of the activity of the pyruvate dehydrogenase complex from beef by and Natl. Acad. Sci. U. S. A. PubMed Scopus Google Scholar). and inactivation of the complex is catalyzed by an which is tightly bound to the and and reactivation are catalyzed by a which is to the complex. It at the time that inactivation of the PDH complex by had not been detected The may in a by after a of our on the and inactivation of the PDH complex. we have been able to pyruvate to be oxidized in we ATP to in the This mechanism was confirmed in the laboratories of S. E. and other with preparations of the PDH complex from other and from pigeon breast In the early the three of enzyme regulation by were and Our results with the PDH complex indicated that this mechanism is more had been in the that in the early was an of system restricted to one limited of with the finding from laboratory that pyruvate dehydrogenase is by the out of the more restricted a period of several years our group separated the and heart PDH complexes into their component enzymes E3, PDH and PDH and characterized the individual enzymes Reed L.J. α-Keto acid dehydrogenase complexes. Purification and properties of the component enzymes of the pyruvate dehydrogenase complexes from and PubMed Scopus Google Scholar). We showed that the undergoes on three We were by the in the electron of negatively stained preparations of the dihydrolipoamide acetyltransferase (E2). subunits appeared to be located at the vertices of a of at the vertices of a our electron studies revealed that are two polyhedral forms of E2, the cube and the (Fig. The a of 24 E2 subunits arranged with symmetry. This is by the E2 components of the E. coli PDH and KGDH complexes and by the E2 components of the KGDH and α-keto acid dehydrogenase complexes. The a of subunits arranged with symmetry. E2 components of this are found in the PDH complexes from and and the A of subunits appeared to be in the of both of polyhedral These were confirmed and extended by results from x-ray diffraction by collaborators David DeRosier and Marvin Hackert and by and The heart PDH complex has a molecular weight of about million. is ∼30 E1 and 12 which are on the E2 by 12 protein (see We proposed that the E1 are located on the 30 edges and the E3 dimers in the 12 faces of the E2 regulation of the PDH complex is fascinating it not only the but also the to the PDH the PDH located in the It is that the major of the activity are and which involve the and respectively. In the early 1970s our group partially purified PDH from heart and and showed that it or for activity. Richard and associates that the activity of the in the presence of and in my group showed that of the to the E2 component of the PDH presumably in to its the of This apparently is the molecular basis of the activation of PDH by other In my laboratory in the early and purified PDH to and showed that it of a and catalytic and a flavoprotein of function later designated showed that the of PDH to Richard and associates showed that the activity of PDH in by the of the to apparently the The function of a in the early and By the properties of and the native PDH bound to we obtained into the function of Reed L.J. of the of pyruvate dehydrogenase Natl. Acad. Sci. U. S. A. PubMed Scopus Google Scholar) found that the of to and that the of PDH but not to apparently by with We these observations to that or the site of and that a in that its These observations the that an on may its of PDH activity. further of in PDH we initiated in the late molecular studies of the PDH complex in the The the five proteins comprising the complex E2, were and Studies on protein confirmed and extended studies of and of and their associates with the protein component of the PDH complex. and E2 apparently from a an lipoyl domain, by an domain, and then by a domain that is involved in to the of E2. The availability of and the provided an to the binding and of and on In the microscopy and electron in with and and their associates revealed a unique structural organization of the complex U. Reed L.J. On the unique structural organization of the pyruvate dehydrogenase complex..J. Biol. Chem. Full Text Full Text PDF PubMed Scopus Google Scholar). As revealed by x-ray and and by electron the of trimers arranged at the vertices of a (Fig. The trimers are by 30 to form an structure, with the of the the center of the Our results showed that the of theE2 trimers within the and an E3 each of the 12 The finding that the of the scaffold the and of binding provides a of the that a of 12 copies of complex. electron microscopy of PDH complex from S. thereof that the of the E2 is also Research on the structural organization of S. PDH complex is electron microscopy studies that individual molecules of an of Reed L.J. evidence for the and of the pyruvate dehydrogenase complex revealed by electron Biol. Chem. Full Text Full Text PDF PubMed Scopus Google Scholar). We have proposed that and of the is and that protein is an component of the function of the PDH multienzyme complex. I these have given of the and I have in this trail of research from lipoic acid to the structure, function, and regulation of the α-keto acid dehydrogenase complexes. I have been in the of this by and and of the of the and by collaborators at other and I to the for Research and the of for R. P. and A. regulation of through pyruvate dehydrogenase and the acid cycle in Soc. S. and C. and protein of the pyruvate dehydrogenase complex of Escherichia coli. Acad. Gunsalus, I. C. transfer and of lipoic acid in The of and University R. and domains in catalytic for multiple P. selection in Soc. L. The and function of lipoic L. complexes. Chem. and M. S. acid dehydrogenase and Acad. and L. of the E2 in the of of pyruvate dehydrogenase complex. M. and R. H. The pyruvate dehydrogenase structure and