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© 2003 by The International Union of Biochemistry and Molecular Biology Printed in U.S.A. BIOCHEMISTRY AND MOLECULAR BIOLOGY EDUCATION Vol. 31, No. 1, pp. 5–15, 2003 Mini-Series: Modern Metabolic Concepts The Biochemistry of the Pyruvate Dehydrogenase Complex* Received for publication, September 5, 2002, and in revised form, September 17, 2002 Mulchand S. Patel‡ and Lioubov G. Korotchkina From the Department of Biochemistry, School of Medicine and Biomedical Sciences, State University of New York at Buffalo, Buffalo, New York 14214 Keywords: Pyruvate dehydrogenase complex, regulation by phosphorylation, diabetes, genetic defects. compound to pyruvate (or its equivalent) for the synthesis of glucose in animals, the flux through PDC is tightly regulated to meet the specific metabolic and energetic needs of different tissues during the fed and fasting (starvation) states. This is accomplished by covalent modification of the ratelimiting component of the complex involving sophisticated interplay among the components of the complex and allosteric modulations by acetyl-CoA and NADH, the products of the reaction (and also of fatty acid oxidation). It is evident from these simple considerations that PDC plays a key role as a gatekeeper of both caloric and glucose homeostasis in mammals. In this review, we will discuss recent developments concerning the structure-function relationship of this multienzyme complex from various organisms with emphasis on regulatory aspects of the mammalian complex. Detailed accounts of various aspects of this complex can be found in several excellent reviews [1–11]. In the Westernized world the daily dietary caloric requirements are roughly provided as follows: 40% from carbohydrates, 40% from fats, and 20% from proteins. In some populations in developing countries the daily caloric contribution from carbohydrate is even higher due to its readily available sources and relatively low cost. Glucose, the principal product of carbohydrate digestion, passes through a series of enzymatic steps first in the non-oxidative glycolytic pathway (from glucose to pyruvate) followed by efficient oxidative metabolism via the tricarboxylic acid cycle (from acetyl-CoA to CO2 and H2O) to harness a portion of its potential energy as ATP. Interestingly, these two major pathways are directly linked by the pyruvate dehydrogenase complex (PDC)1 localized in the mitochondrial matrix (Fig. 1). In the fed state acetyl-CoA generated from pyruvate (derived mostly from glucose and some dietary amino acids) is also utilized for the biosynthesis of lipids such as long chain fatty acids and cholesterol by lipogenic tissues (such as liver and adipose tissues and under special conditions in mammary glands during lactation and in the brain during the prenatal and early postnatal development). Additionally, amino acids from excess dietary protein are metabolized by several specialized reactions or pathways generating intermediates that have to be ultimately converted to pyruvate first and then to acetylCoA via PDC either for complete oxidation to CO2 and H2O or for lipogenesis. PDC is the only known reaction in most eukaryotes to generate acetyl-CoA (two-carbon compound) from pyruvate (three-carbon compound). Since this is a physiologically irreversible reaction and since there is no other known reaction or pathway to convert the two-carbon STRUCTURE AND ORGANIZATION OF PDC PDC is present in most prokaryotic and eukaryotic organisms. It catalyzes several sequential reactions of oxidative decarboxylation of pyruvic acid by the action of its three catalytic components: (i) pyruvate dehydrogenase (E1) catalyzing the decarboxylation of pyruvate followed by reductive acetylation of lipoyl moieties covalently linked to the dihydrolipoamide acetyltransferase (E2), the second catalytic component of the complex; (ii) E2 catalyzing the formation of acetyl-CoA; and (iii) dihydrolipoamide dehydrogenase (E3) reoxidizing the reduced lipoyl moieties of E2 with the consequent reduction of NAD⫹ to NADH (Fig. 2) [12]. PDC is a multienzyme complex with a well organized structure that has two different morphologies based on symmetry of the central E2 core, i.e. octahedral for Gram-negative bacteria and icosahedral for eukaryotes (mammals, yeast, plants, and nematodes) and some Gram-positive bacteria. E2 consists of three well defined domains connected by flexible linkers: (i) inner domains interacting noncovalently with each other to form the core of the complex and each having the catalytic site; (ii) subunit-binding domain interacting with E1 and E3 (for PDC from bacteria, plants, and nematodes); and (iii) the lipoyl domain (containing up to three repeating units, one for yeast and Gram-positive bacteria, two for mammals, and three for Gram-negative bacteria; Fig. 3) providing coupling of the E1, E2, and E3 active sites by transferring * This work was supported by United States Public Health Service Grants DK20478 and DK42885. ‡ To whom correspondence should be addressed: Dept. of Biochemistry, School of Medicine and Biomedical Sciences, State University of New York at Buffalo, 140 Farber Hall, 3435 Main St., Buffalo, NY 14214. Tel.: 716-829-3074; Fax: 716-8292725; E-mail: [email protected]. 1 The abbreviations used are: PDC, pyruvate dehydrogenase complex; E1, pyruvate dehydrogenase; E2, dihydrolipoamide acetyltransferase; E3, dihydrolipoamide dehydrogenase; BP, E3binding protein; PDK, pyruvate dehydrogenase kinase; PDP, phosphopyruvate dehydrogenase phosphatase; TPP, thiamine pyrophosphate; PPAR␣, peroxisome proliferator-activated receptor ␣; CRE, cAMP-response element; CAT, chloramphenicol acetyltransferase; MEP, mouse E1␣ promoter site. This paper is available on line at http://www.bambed.org 5 6 BAMBED, Vol. 31, No. 1, pp. 5–15, 2003 FIG. 1. Regulation of PDC activity by phosphorylation/dephosphorylation catalyzed by PDKs and PDPs. Three serine residues of E1 phosphorylation sites are shown as S1, S2, and S3 in dephosphorylated condition and as P1, P2, and P3 when phosphorylated. FIG. 2. Overall PDC reaction. CH3-COH⫽TPP, 2-␣-hydroxyethylidene-TPP; Lip-S2, oxidized lipoyl moiety of E2. an acetyl group and reducing equivalents. The core of Gram-negative bacterial PDC is composed of inner domains of 24 subunits of E2 and binds 24 dimers of E1(␣2) and 12 dimers of E3(␣2) (Table I). PDCs with icosahedral symmetry have 60-mer E2 cores that bind up to 30 tetramers of E1(␣22) and 12 dimers of E3. Eukaryotic organisms have an additional component in PDC not present in bacterial complexes, i.e. E3-binding protein (BP) [6, 12]. BP has a domain structure similar to E2 structure except: (i) only one lipoyl domain is present; (ii) the subunit-binding domain binds E3 and not E1; and (iii) the inner domain does not perform a catalytic function (due to the absence of the critical histidine residue). The PDC core of eukaryotes is composed of the inner domains of E2 and BP [13]. Ascaris suum PDC has a BP without a lipoyl domain and only in the anaerobic form of this nematode [14]. Mammalian, plant, and nematode PDCs are regulated by reversible phosphorylation/dephosphorylation that requires additional regulatory enzymes, pyruvate dehydrogenase kinase (PDK), and phosphopyruvate dehydrogenase phosphatase (PDP) (Table I). The three-dimensional structures of several components of PDC were determined by x-ray crystallography: E1 from Escherichia coli [15] and human2; E1 subunit from Pyrobaculum aerophilum [6]; cubic core of E2 (consisting of the inner domains) from Azotobacter vinelandii, Bacillus stearothermophilus, and Enterococcus faecalis; subunit-binding domain of E2 from B. stearothermophilus; E2 lipoyl domains from B. stearothermophilus, E. coli, and A. vinelandii; and E3 from yeast, A. vinelandii, Pseudomonas putida, and Pseudomonas fluorescens [5, 6, 16]. So far it is not possible to solve the three-dimensional structure of the whole PDC because of its huge molecular mass of 8 –10 million Da, hollow structure of its inner core with high solvent content, and flexibility of E2 domains. However, the structures of the E2 core and entire PDC from yeast and mammals were studied by cryoelectron microscopy (Fig. 4) [17, 18]. Both electron microscopy and crystal structures of the E2 core revealed the trimer building blocks in the cubic (8 trimers) or dodecahedral (20 trimers) structures. The trimers are connected by bridges and form a structure with large openings and a cavity inside. Substrates of E2 enter the active site from different directions, i.e. CoA come from inside the core, while lipoyl moieties come from the outside. Twelve monomers of BP (for eukaryotic PDCs) were suggested to bind within the cavity of E2 and interact with the E3 dimer inside 12 pentagonal openings. Yeast E2 core has the size of 250 Å with 40-Å variations in diameter as found by electron microscopy. The contraction and expansion of the E2 core was proposed to reflect the conformational mobility of PDC during 2 E. Ciszak, L. G. Korotchkina, P. Dominiak, S. Sidhu, and M. S. Patel, unpublished observations. 7 FIG. 3. The structural domains of E2 and BP. TABLE I Composition of the pyruvate dehydrogenase complex from different species Oligomeric structure Source Isoenzymes ␣2 ␣2 ␣22 ␣22 ␣22 ␣22 ␣22 E. coli A. vinelandii B. stearothermophilus Mammals Nematode Yeast Plant ␣ somatic/␣ testis E1-I/E1-II E2 ␣24 (65–66 kDa) ␣60 (49–76 kDa) ␣24 E3 (49–58 kDa) ␣2 BP (45–48 kDa) ␣ PDK (39–48 kDa) ␣2 E. coli A. vinelandii B. stearothermophilus Mammals Nematode Yeast E. coli A. vinelandii B. stearothermophilus Mammals Nematode Yeast Plant Mammals Yeast Nematode Mammals Components E1 ␣2 (89–99 kDa) ␣22 ␣ (40–43 kDa)  (35–37 kDa) ␣60 ␣2 ␣2 PDP Ca (52 kDa) Rb (96 kDa) a b C, catalytic subunit. R, regulatory subunit. C/R Nematode Plant (Arabidopsis thaliana) (maize) Mammals Mitochondrial/ plastid PDK1 PDK2 PDK3 PDK4 PDK PDK1/PDK2 C: PDP1/PDP2 8 BAMBED, Vol. 31, No. 1, pp. 5–15, 2003 FIG. 4. Three-dimensional reconstruction of bovine PDC (A and B) and diagrammatic representation of the structural domains of E2 subunit (C). Shaded-surface representation of 3-fold axes of symmetry of the three-dimensional reconstruction of the bovine kidney PDC (A) and with the closest half removed to reveal the linker (blue) that binds E1 (yellow) to the E2 core (green; B). The inner linker is 50 Å in length. C, the C-terminal self-association domain is responsible for the assembly of the dodecahedral scaffold to which E1 and BP䡠E3 bind. The N-terminal half of the E2 comprises the L1 and L2 lipoyl domains, the E1-binding domain, and their associated linkers. The inner linker is revealed in the three-dimensional structure of the PDC. 5-, 3-, and 2-fold axes are indicated. Reprinted with permission from Zhou et al. [18]. Copyright 2001 National Academy of Sciences, U. S. A. catalysis [17]. Recently the structure of the whole PDC from bovine kidney was constructed based on cryoelectron microscopy (Fig. 4) [18]. Bovine PDC was found to have 60 monomers of E2, 12 monomers of BP, 22 tetramers of E1, and 6 dimers of E3. The size of the complex in the presence of E1 molecules is about 500 Å. E1 molecules are organized in trimers bound through the subunitbinding domains of E2 70 Å above the trimers of inner domains of E2. Lipoyl domains are proposed to rotate around the subunit-binding domains transferring intermediates between active sites of E1, E2, and E3, and this movement is provided by the conformational changes of the E2 core. E1 of PDC is a dimer of identical subunits in Gramnegative bacteria or a heterotetramer composed of 2 ␣ and 2  subunits in eukaryotes and Gram-positive bacteria. Both types of E1 have two active sites and require thiamine pyrophosphate (TPP) and Mg2⫹ as cofactors. Sequence similarity between the two types of E1 is low; however, there is a significant similarity in the structure of the TPP-binding region for several TPP-dependent enzymes, the three-dimensional structures of which are known (yeast transketolase, Lactobacillus plantarum pyruvate oxidase, yeast, and Zymomonas mobilis pyruvate decarboxylase, P. putida benzoylformate decarboxylase, human and P. putida branched-chain ␣-keto acid dehydrogenase, Desulfovibrio africanus pyruvate:ferredoxin oxidoreductase [19, 20], and E. coli PDC-E1 [15]). The common features of the active sites of these proteins are: (i) active sites are localized on the interface of two subunits; (ii) two residues of the TPP motif (a sequence of 30 –33 amino acid residues conserved for all known TPP-requiring enzymes) coordinate binding of the pyrophosphate moiety of TPP through interactions with a divalent cation (Mg2⫹ or Ca2⫹); (iii) amino acid residues help TPP to maintain the “V” conformation necessary for catalysis (among them are phenylalanine or tyrosine residues providing a stacking interaction with the aminopyrimidine ring of TPP and a hydrophobic residue between two aminopyrimidine and thiazole rings); and (iv) N1⬘ of the pyrimidine ring binds to the glutamine residue through a hydrogen bond, which affects reactivity of the 4⬘-amino group of TPP and activates the C2 hydrogen of the thiazolium ring of TPP. Only the E. coli PDC-E1 structure has been reported so far. The human PDC-E1 structure was solved recently.2 The structure-function relationships of E1 from human and E. coli have been studied intensively using site-directed mutagenesis [21–23]. Mammalian E1 has two isoforms (with 92% homology), a somatic cell form with the gene (PDHA-1 for human) located on the X chromosome and a testis-specific form with the gene (PDHA-2 for human) on chromosome 4 [24]. The second isoform is necessary during spermatogenesis when the X chromosome is inactivated or absent. A. suum has two E1 isoforms, isoform I in anaerobic adults and isoform II in aerobic larvae [25]. E3 is a homodimer with two active sites localized at the dimer interface. Each active site contains tightly but noncovalently bound FAD, which participates in the electron transfer from dihydrolipoamide to NAD⫹ with participation of an active site disulfide. Each subunit of E3 has four structural domains: FAD-binding domain, NAD⫹-binding domain, central domain, and interface domain. Interestingly, E3, the product of the same gene, participates in four different complexes, i.e. PDC, ␣-ketoglutarate dehydrogenase complex, branched-chain ␣-keto acid dehydrogenase complex, and as H-protein in glycine synthase. Only in P. putida do three different genes code for three dihydrolipoamide dehydrogenases. E3 is bound to the subunit-binding domain of BP in eukaryotic PDC and to the subunit-binding domain of E2 in other complexes. The structure of B. stearothermophilus E3 bound to the subunit-binding domain of E2 was solved to 2.6-Å resolution [16]. The interaction between these proteins was shown to be electrostatic, i.e. between positively charged residues of the binding domain and negatively charged residues of E3. The same subunit-binding domain of E2, however, 9 binds E1 at the different site. While E3 interacts with the N-terminal part of the subunit-binding domain, E1 binds to the C-terminal part of the same domain. The binding site for the A. vinelandii E1 was shown to involve not only the subunit-binding domain but also the E2 inner domain [9]. PDK is a specific kinase that phosphorylates E1 of PDC and is present in human, rodent, plant, nematode, and fruit fly. PDK has several isoforms, four in mammalian PDC (PDK1, PDK2, PDK3, and PDK4), two in plants, and one in nematode and fruit fly (Table I) (see Ref. 26). PDK, a serine-specific kinase, phosphorylates three specific serine residues named site 1, site 2, and site 3 based on the rate of phosphorylation of mammalian E1 [27], isoform II of A. suum E1, and fruit fly E1. Only two phosphorylation sites are present in isoform I of A. suum E1 and in Caenorhabditis elegans, and only one site is in plant E1s. Yeast E1 has phosphorylation site 1 in a sequence that can be phosphorylated by bovine PDK; however, PDK is not detected in yeast. Mammalian PDKs have low homology with eukaryotic serine protein kinases and are more related to bacterial histidine protein kinases [28]. Recently the threedimensional structure of rat PDK2 was solved [29]. The C-terminal domain of PDK2 appeared to be similar to the nucleotide-binding domain of bacterial histidine kinases. However, PDK catalyzes the phosphate transfer directly to the serine residue of the substrate and not through a histidine residue of the kinase itself as bacterial histidine kinases do, and hence PDK is related to the eukaryotic serine kinases and ATPases by catalytic mechanism. Each PDK isoenzyme is a dimer composed of identical subunits. PDKs were shown to bind the lipoyl domains of E2 and BP [6]. In mammalian PDC there are two lipoyl domains in E2 (L1 and L2) and one in BP (L3) (Fig. 3). Different isoenzymes of PDK have preference to the lipoyl domains: PDK1 binds to L1 and L2; PDK2 and PDK3 prefer L2 but can also bind L1 less efficiently; and PDK4 binds to L3 and less to L1 [6]. The number of PDK molecules in the PDC is as low as one to three molecules per complex. Transfer of PDK from one lipoyl domain to another is necessary to achieve phosphorylation of multiple E1 molecules. Moreover, PDK is activated when bound to the lipoyl domain of E2 (the degree of activation is different for different isoenzymes of PDK). The activation may be achieved by the conformational changes of PDK when bound to the lipoyl domain and co-localization of PDK and its substrate E1 on E2 because activation by a free lipoyl domain is less compared with E2. E2 does not only bind and transfer PDK in PDC but also regulates the activity of PDK through the state of the lipoyl groups. PDK activity is stimulated with the reduction and reduction/acetylation of the lipoyl groups of E2 occurring during PDC reaction [6]. The stimulation is explained by an allosteric effect of a reduced or acetylated form of lipoate of the lipoyl domain involved in the PDK binding. The extent of stimulation depends on the PDK isoenzyme with the maximum stimulation found for PDK2. PDP is a heterodimer composed of two nonidentical subunits, the catalytic subunit belonging to the phosphatase 2C class of phosphatases and the regulatory subunit, which is a flavoprotein (Table I). Two isoenzymes of the catalytic subunit of PDP are identified in mammalian tis- TABLE II Tissue distribution of mRNAs of mammalian PDK isoenzymes Tissue distributions are based on citations in Ref. 7. Tissue PDK1 Heart Skeletal muscle Pancreas Liver Kidney Testis Brain Spleen Lung ⫹⫹⫹⫹⫹ ⫹⫹⫹ ⫹ ⫹⫹ PDK2 ⫹⫹⫹⫹ ⫹⫹⫹⫹ ⫹⫹⫹ ⫹⫹⫹ ⫹⫹⫹ ⫹⫹⫹ ⫹⫹⫹ ⫹ ⫹ PDK3 PDK4 ⫹ ⫹⫹⫹⫹ ⫹⫹⫹⫹⫹ ⫹⫹ ⫹⫹⫹⫹ ⫹⫹ ⫹ ⫹⫹⫹ ⫹⫹ ⫹ ⫹⫹⫹ sues (PDP1 and PDP2) with different tissue distribution, activities, and regulation [7]. The regulatory subunit of PDP1 affects the sensitivity of the catalytic subunit to Mg2⫹. Mammalian PDP1 binds to the L2 of E2 with Ca2⫹ playing a bridging role [6]. REGULATION OF PDC Short-term Regulation of PDC Activity—The central role of PDC in glucose homeostasis necessitates the high level of regulation of its activity in mammalian tissues. Regulation of PDC activity within minutes and hours will be referred to as a short-term regulation, while regulation exerted at a transcriptional level within days to weeks is considered as a long-term regulation. The short-term regulation of PDC by metabolite effectors and hormones is achieved through phosphorylation of E1 by PDK, leading to inactivation of E1 (and hence PDC), and dephosphorylation and reactivation is accomplished by PDP (Fig. 1). As discussed above four isoenzymes of PDK and two isoenzymes of the catalytic subunit of PDP are present in mammalian tissues. Isoenzymes of PDK and PDP have different specific activities, sensitivity to different effectors, and tissue-specific distribution (Table II) [6, 7]. The highest amount of PDK1 mRNA is present in heart, and decreasing levels are present in skeletal muscle, liver, and pancreas. PDK2 is expressed in many tissues with low amounts in spleen and lung. PDK3 mRNA is maximal in testis and less in lung, brain, and kidney. PDK4 mRNA was detected in skeletal muscle and heart with low levels in lung, liver, and kidney. A high level of PDP1 is present in skeletal muscle, while levels of PDP2 are higher in liver and adipose tissue [7]. The presence of several phosphorylation sites of E1 adds to the complexity of the mechanism of regulation of PDC activity. In mammalian PDC three phosphorylation sites are phosphorylated differently by different isoenzymes of PDK [26]. PDK1 phosphorylates all three sites, whereas PDK2, PDK3, and PDK4 can modify only sites 1 and 2 when PDKs are bound to E1 in PDC. The activities of PDKs toward site 1 are: PDK2 ⬎ PDK4 ⬎ PDK1 ⬎ PDK3; the activities for site 2 are: PDK3 ⬎ PDK4 ⬎ PDK2 ⬎ PDK1. In the absence of E2 PDKs are also active. The maximum activity in the absence of E2 was demonstrated toward site 1; however, site 2 could be phosphorylated by PDK4, and site 3 could be phosphorylated by PDK1 in the absence of E2. Phosphorylation of each site results in E1 inactivation. The mechanism of inactivation of E1 by phosphorylation of three sites is site-specific [30]. Modification 10 of site 1 prevents interaction of the E1 active site with the substrates of the reaction, pyruvate and more with the lipoyl domain of E2, whereas phosphorylation of site 3 affects E1 interaction with TPP. The products of the PDC reaction (and more importantly the products of fatty acid and ketone body oxidation), NADH and acetyl-CoA, increase activity of PDK through reduction/acetylation of lipoyl moieties of the lipoyl domains of E2 to which PDKs are bound. Hence the NADH/ NAD⫹ and acetyl-CoA/CoA ratios determine the portion of PDC present in active (dephosphorylated) form. The highest stimulation is achieved for PDK2. The least sensitive isoenzyme is PDK3 [6, 26]. Activities of PDKs also depend on the ATP/ADP ratio as ADP and phosphate anion are inhibitors of PDK activity. PDK2 and to a lesser extent other PDK isoenzymes are inhibited by pyruvate, the substrate of the PDC reaction [6]. Inhibition by pyruvate is synergistic with ADP inhibition. The mechanism of inhibition was suggested to be formation of the dead end complex PDK-ADP-pyruvate that prevents ADP dissociation and hence PDK activity [6]. Activities of PDP isoenzymes are regulated by the concentrations of divalent metal ions Mg2⫹ and Ca2⫹ [6]. PDP is a Mg2⫹-dependent enzyme, and hence changes in Mg2⫹ concentration along with the ATP concentration in the cell affect PDP activity. PDP2 is less sensitive to Mg2⫹ concentration than is PDP1. Sensitivity of PDP1 to Mg2⫹ concentration changes when it is bound to the regulatory subunit of PDP. Polyamines (spermine) increase the activities of both PDP2 present alone and PDP1 associated with a regulatory subunit. Ca2⫹ ions (increased in conditions of energy deficit) elevate activity of PDP1 by facilitating its binding to E2. The short-term hormonal regulation of PDC is achieved by insulin, which increases activity of PDP resulting in dephosphorylation and activation of PDC. The signal from insulin is transferred by protein kinase C␦ that translocates to mitochondria where it phosphorylates and activates PDP [31]. Plants have both mitochondrial and plastid PDC, and only mitochondrial PDC is regulated by phosphorylation [32]. Interestingly, activity of PDC in plants depends on light. Photosynthesis and photorespiration cause a decrease in mitochondrial PDC activity due to phosphorylation by the elevated amount of ATP. In contrast, the increase of pH and Mg2⫹ concentration in plastid in light during photosynthesis enhances plastid PDC activity. Developmental conditions of nematodes are reflected in the regulation of PDC. Nematodes have two isoforms of E1, E1-I present in anaerobic adult muscle and E1-II in aerobic larvae [25]. The two E1 isoforms have different stoichiometry of phosphorylation and inactivation. Three phosphorylation sites in E1-II isoform are modified similarly to mammalian E1. Anaerobic E1-I isoform has only two phosphorylation sites that are more resistant to inactivation. This fact together with the reduced sensitivity of nematode PDK to the elevated NADH/NAD⫹ and acetyl-CoA/CoA assist in maintaining the PDC activity at a high level in anaerobic conditions when in the absence of the tricarboxylic acid cycle the products of the PDC reaction are used for fatty acid biosynthesis. One wonders why it is critical to have such a tight and BAMBED, Vol. 31, No. 1, pp. 5–15, 2003 extensive regulation of PDC function with so many phosphorylation sites as well as multiple isoforms of kinases and phosphatases that are regulated by such a variety of control factors. This may at first seem like overkill in regulation of PDC! Based on the available information, the following explanations are considered for such an elaborate regulation of PDC. First, during the transition from the fed to fasting state, the organism needs to conserve the three-carbon compounds (such as pyruvate, lactate, and alanine) from further degradation so that they can be used for gluconeogenesis (Fig. 5). The phosphorylation mechanism renders the complex completely inactive by phosphorylating even only one of the three phosphorylation sites present in the ␣ subunit of E1 (two ␣ subunits present in E1 provide six potential sites for phosphorylation, and phosphorylation of any one site is sufficient to inactive E1 and hence PDC). There are two possible reasons for the multiple sites of phosphorylation: (i) the gradual reactivation by dephosphorylation of multisite phosphorylated E1 by the phosphatases during the conditions such as the transition from the fasted to fed state and (ii) providing a fall back control to continue to maintain this crucial regulation if one phosphorylation site is modified by genetic mutation. Second, the importance of the presence of multiple forms of specific kinases with differences in their regulation is linked to their tissue-specific distribution. Since all tissues are not created equal in their roles in maintaining glucose homeostasis, the distribution of kinases in different tissues (such as increases in PDK2 and PDK4 activities in liver during fasting and in diabetes) plays critical roles in regulating the carbon flux through PDC. Additionally, the sensitivity of PDKs to the changes in the NADH/NAD⫹, acetyl-CoA/CoA, and ATP/ADP ratios due to increases in the oxidation of fatty acids (and ketone bodies in extrahepatic tissues after an extended fasting period) not only speeds up the action of specific kinases but also maintains control over an extended period. Third, the changes in metabolites such as NADH, acetylCoA, and ATP in the mitochondria due to increased oxidation of fatty acids, an alternate fuel source for acetylCoA, result in drastic changes in the ratios of NADH/ NAD⫹, acetyl-CoA/CoA, and ATP/ADP in the mitochondria that allosterically stimulate the activities of most PDKs. Although the same metabolite changes also regulate the activity of PDC at one of its component enzymes, the degree of inhibition exerted is generally not sufficient to shut off the complex; whereas the very same changes in the metabolite levels through the allosteric activation of PDKs amplify the process of inactivation! This type of amplification control is linked to the availability of other oxidizable fuels in the mitochondria of different tissues. For example, the levels of the active state of PDC in the liver and skeletal muscles are rapidly decreased during the fed to fasting transition, hence enabling these tissues to oxidize fatty acids and sparing complete oxidation of glucose-derived three-carbon metabolites (Fig. 5). In contrast, the levels of the active state of PDC in the brain is maintained fairly constant during the fed-fasting-refeeding transitions under normal dietary conditions (meaning the capacity of the brain to oxidize glucose to CO2 and water 11 FIG. 5. Changes in plasma hormonal levels, intramitochondrial metabolites, and glucose metabolism in mammalian liver, skeletal muscle, and brain during the fed and fasted states. The central role of PDC in glucose disposal during the fed state and in glucose sparing during the fasted state is highlighted. The relative flux of glucose, fatty acids (FAs), and ketone bodies in the liver, skeletal muscle, and brain are shown by heavy and thin arrows. TCA, tricarboxylic acid cycle; VLDL, very low density lipoproteins; OAA, oxaloacetate; PEP, phosphoenolpyruvate; AAs, gluconeogenic amino acids; I, insulin; G, glucagon. is not affected during an overnight fast or short fasts of 1–2-day duration) (Fig. 5). This allows the brain to continue uninterrupted oxidation of glucose for energy production. However, when the plasma concentrations of ketone bodies are sufficiently increased over an extended period of fasting (early starvation period) or in uncontrolled diabetes, the brain is gradually able to oxidize increasing amounts of ketone bodies. That generates the metabolites (such as acetyl-CoA and NADH) that reduce PDC activity via their actions on PDKs and hence spare glucose oxidation while maintaining energy homeostasis in the brain. Fourth, the general structural and catalytic similarities of PDC with the other two related complexes in the mitochondria such as the ␣-ketoglutarate dehydrogenase complex and the branched-chain ␣-keto acid dehydrogenase complex shed light on the need for regulation or an absence of regulation of these complexes based on their importance in the oxidative metabolism. For example, there is a complete lack of regulation by phosphorylation/ dephosphorylation of the ␣-ketoglutarate dehydrogenase complex because of its central location in functioning of the tricarboxylic acid cycle. Since the tricarboxylic acid cycle is involved in the oxidation of all metabolites gener- ated from the initial metabolism of carbohydrates, fats, and amino acids, which are capable of feeding into the cycle for further oxidation, the elaborate regulation of this complex would not provide any advantage for fuel-specific switching in the tissue. In contrast, the regulation of the branched-chain ␣-keto acid dehydrogenase complex by a phosphorylation/dephosphorylation mechanism does provide a control for oxidation of the three essential amino acids (leucine, isoleucine, and valine), which play a pivotal role in nitrogen metabolism in the body. Thus, it is no surprise that this complex is regulated by phosphorylation of one functional site only on its E1 component by a specific kinase and a specific phosphatase with only a limited effect, if any, of metabolite-mediated control of this process. Hence, compared with the elaborate control scheme of regulation of PDC, the branched-chain ␣-keto acid dehydrogenase complex is controlled by a less elaborate system for its regulation and may reflect a limited, specific role of this complex in the scheme of overall metabolism for energy homeostasis. Long-term Regulation of PDC Activity—Although the short-term regulation of PDC by covalent modification is the key to the maintenance of glucose homeostasis during 12 BAMBED, Vol. 31, No. 1, pp. 5–15, 2003 FIG. 6. Schematic presentation of specific cis-acting elements and nuclear protein-binding domains of the proximal promoter regions of the human PDHA-1 and PDHA-2 genes. ChoRE, carbohydrate-response element. the transition between the fed and fasted states, alterations in total PDC activity (and hence PDC content) in response to dietary and hormonal changes over longer periods (long-term regulation) are also observed. For instance, total PDC activity in liver and adipose tissue increased by ⬃1.5- and 3.5-fold, respectively, in rats fed a high sucrose diet for 1–2 weeks compared with that of chow-fed controls [33]. The levels of the component proteins of PDC measured by immunological analysis in livers from high sucrose-fed animals closely correlated with changes in total PDC activity [34]. In contrast, total PDC activity was significantly decreased in livers from high fat-fed rats. Interestingly, total and active PDC levels were not affected in heart and skeletal muscle of rats fed the high sucrose or high fat diets. Hypothyroidism in the rat caused about one-third reduction in total PDC activity in the liver, and this change was correlated with a similar reduction in amounts of E1 proteins (the other components of PDC were not measured) [33]. These stable changes in PDC activity are explained by long-term regulation occurring at the transcriptional level. Several genes encoding for PDC components were shown to be regulated through different cis-acting elements in their promoter regions [4]. Interestingly, culturing of hepatoma HepG2 and insulinoma MIN6 cells in medium containing high glucose levels resulted in increased total PDC activity, and this increase was correlated with the levels of E1␣ and E1 mRNA levels [35]. The elevated glucose presumably increases transcription of E1␣ and E1 in situ due to the presence of carbohydrate-response elements in the promoters of the E1␣ and E1 genes [35]. The major determinant of the PDC activity under different physiological conditions is the amount of PDK. An increase in the expression of PDK isoforms in different pathophysiological conditions is well documented and intensively studied [7]. Of the four isoenzymes of PDK, the highest degree of regulation is observed for PDK4 and to a lesser extent for PDK2. However, the abundance of PDK2 in all tissues makes its regulation very important. Starvation increases the amounts of PDK2 and PDK4 in liver, kidney, lactating mammary gland, and fast-twitch muscle and PDK4 levels in heart and slow-twitch muscle [7]. During starvation, mRNA, but not the protein of PDK4, is higher in white adipose tissue and brain. Brain is the only tissue in which PDC activity does not change during fasting because brain depends on glucose as a major source of energy. Diabetes increases the expression of PDK4 in heart and skeletal muscle and slightly increases the expression of PDK2 in skeletal muscle [7]. Insulin treatment of diabetic animals rapidly reverses the changes in PDK4 activity [7]. In adipose tissue PDC activity is highly regulated by insulin, which activates PDP after refeeding of starved animals or by administration of insulin to diabetic animals. Hyperthyroidism results in increased amounts of PDK4 protein in heart and muscle and of PDK2 protein in fasttwitch muscle [36]. Starvation, diabetes, and a high fat diet are conditions leading to higher levels of free fatty acids. Fatty acids are known to activate peroxisome proliferatoractivated receptor ␣ (PPAR␣). An agonist of PPAR␣ (WY14,643) is found to up-regulate expression of PDK4 in rat skeletal muscle and liver and in cultured hepatoma cells [7, 11]. Glucocorticoid (dexamethasone) causes the same response in hepatoma cells. Both compounds increased PDK4 mRNA without altering its stability. Insulin, on the other hand, diminishes the levels of PDK4 and PDK2 mRNAs. It is proposed that elevated levels of fatty acids in certain physiological conditions increase expression of PDK4 and PDK2 through activation of PPAR␣. The expression of PDK4 and PDK2 is suggested to be regulated to some extent by the drop in insulin [37]. Recently expression of PDP2 was reported to decrease in diabetes and starvation, resulting in lower rates of dephosphorylation [38]. Up-regulation of PDK expression and down-regulation of PDP expression leads to hyperphosphorylation (phosphorylation on multiple sites) of E1 and inactivation of 13 PDC. Phosphorylation of any one of the E1 phosphorylation sites (site 1 is modified first in the cell) is sufficient for inactivation; however, hyperphosphorylation results in a longer duration of inactivation. Complete dephosphorylation of more than one site requires a longer period for reactivation after changing physiological conditions. REGULATION OF PDC GENES Gene organization and functional analysis of the promoter-regulatory regions of the three human PDC genes (E1␣, E1, and E3) and the mouse Pdha-2 gene have been reported [33]. Two genes have been found to encode the ␣ subunit of the E1 component of PDC in most mammals. Human PDHA-1 and rodent Pdha-1, both localized on chromosome X, are expressed in somatic cells, whereas human PDHA-2 (on chromosome 4) and rodent Pdha-2 (on chromosome 19, band B in the mouse) are expressed in testis only [24]. Of interest are the promoters of the PDHA-1 and PDHA-2 (and Pdha-2 of mouse) genes (Fig. 6). The presence of several cis-acting elements as well as seven distinct nuclear protein-binding domains in the ⫺763/⫹33 promoter region of the human PDHA-1 gene have been identified by sequence analysis and DNase footprinting analysis, respectively [39]. A detailed mutational analysis of the ⫺100 bp promoter region of the human PDHA-1 gene indicated that the overlapping CRE/ TATA-like sequence (⫺24 to ⫺34 bp) functioned as the CRE-like sequence and had a minimal affinity for the TATA-binding protein, indicating this promoter functions as a TATA-less promoter. Furthermore, it was observed that the CRE site is functionally important, and the GCGG sequence at the ⫺52 to ⫺49 bp region was indispensable for the minimal promoter activity.3 A transgene containing ⫺763/⫹33 bp of the human PDHA-1 promoter/structural chloramphenicol acetyltransferase (CAT) gene was responsive to diet-induced expression of CAT activity in kidney and adipose tissue but not in the brain of transgenic mice [40]. However, this transgene construct did not contain sufficient sequence information to direct regulated CAT expression in several tissues. Subsequently it was shown that the ⫺1.7/⫺2.2 and ⫺5.2/⫺5.9 kb regions of the PDHA-1 promoter may possess negative regulatory elements that are likely to function in a tissue-specific manner. In another study, the ⫺100 bp promoter region of the human PDHA-1 gene was shown to be responsive to glucose in transiently transfected HepG2 cells. Mutational analysis of the human PDHA-1 gene promoter region uncovered the presence of two sequences from ⫺78 to ⫺73 bp (CCCCTG) and from ⫺8 to ⫺3 bp (GCGGTG) that are responsible for the glucose-induced increases in promoter activity [35]. Of these two sequences representing new variations of the carbohydrate-response element identified in other glucose-regulated genes, the former sequence (CCCCTG) exhibited a large effect on promoter activity. It should be noted that glucose-mediated stimulation of human PDHA-1 promoter was relatively low (3–5-fold only) compared with that of the L-pyruvate kinase gene and the S14 gene. This could be due to the lack of two copies of 3 J. Tan, R. Dey, H.-S. Yang, and M. S. Patel, unpublished observations. the perfect CACGTG sequence separated by an ideal fivenucleotide spacer in the human PDHA-1 promoter [35]. In contrast to regulation of expression of the PDHA-1 gene promoter, expression of Pdha-2 in mice is tightly regulated so that its transcription occurs only during specific stages of spermatogenesis [24, 41]. Extensive analysis of a core mouse Pdha-2 promoter region has revealed four regions of protection. Two of these regions contain potential motifs for Sp1 and activating transcription factor/ cAMP-response element-binding protein (ATF/CREB) and the remaining two binding sites harbor novel cis-motifs designated MEP-2 (mouse E1␣ promoter site 2) and MEP-3. MEP-2 forms a complex with a putative testisspecific binding factor (-MEP-2BF) [41]. Analysis of the proximal promoter of human PDHA-2 gene showed the presence of four testicular nuclear protein-binding domains within the ⫺106/⫹9 region and three cis-acting motifs, namely MEP-2, Sp1, and Ets (Fig. 6). Functional studies also identified the presence of both enhancer and repressor elements (Ets) in the PDHA-2 promoter that are only expressed in mature sperm [42]. MOLECULAR DEFECTS IN PDC COMPONENTS PDC deficiency is one of the major genetic disorders resulting in congenital lactic acidosis [43]. More than 150 cases of PDC deficiency have been reported with heterogeneous clinical manifestations that are limited largely to the central nervous system due to the brain’s dependence primarily on glucose oxidation for energy production [4, 9, 43, 44]. Given that the human PDC is composed of several different gene products, a genetic defect involving any one of them (except PDKs) could lead to the impairment in its function. Approximately 80% of all reported cases of PDC deficiency involve defects of the E1 component, all of which are found in the ␣ subunit, and the remainder are distributed among the other components except PDKs. A PDK deficiency would not cause lactic acidosis, and the presence of four isoenzymes with overlapping tissue distribution may compensate for deficiency of one isoform. Perhaps only patients with partial defects in which there is some residual PDC activity can survive. This point is well illustrated by null mutation of the Pdha-1 gene in mice using homologous recombination techniques resulting in embryonic lethality [45]. Since the human PDHA-1 gene is localized on chromosome X, the pattern of inactivation of chromosome X in different cells and tissues in affected female patients contributes to the observed variability in residual PDC activity with associated clinical manifestations. In summary, PDC deficiencies further illustrate the importance of PDC in glucose metabolism in the developing mammalian brain. A large number of specific mutations associated with the PDC components have been identified, almost all of which are within the E1␣ coding region [4, 9, 43, 44]. Interestingly, about one-half of E1␣ mutations result from missense codon changes preserving the primary structure and some residual catalytic function of the protein. Several of these missense mutations (at Met-181, Pro-188, Arg349, His-15, and Arg-234, based on numbering of the residues in mature ␣ subunit protein) were recreated by overexpressing human mutant E1 in E. coli to characterize 14 BAMBED, Vol. 31, No. 1, pp. 5–15, 2003 the biochemical basis for altered function [21, 22]. Interestingly, many of these mutations alter the binding of TPP in the E1 active site. To minimize recurrence of lactic acidosis, rational treatments have included the use of ketogenic diets (high in fat content and extremely low or even absent in carbohydrates in a diet), daily administration of large amounts of thiamine (to increase intramitochondrial TPP levels for some mutant proteins), and administration of dichloroacetate (to inhibit PDK and possibly increase the residual PDC in a more active state) [43]. Unfortunately, all of these interventions have met with variable and very limited successful clinical outcomes. CONCLUSION The central role of PDC in glucose homeostasis has been recognized for several decades and has been emphasized in textbooks of biochemistry as a pacesetter for glucose metabolism during fed/fasting/refeeding transitions normally occurring during the 24-h period. The carbon flux through PDC is meticulously controlled by an elaborate mechanism via phosphorylation/dephosphorylation that is not only extremely sensitive to intramitochondrial metabolic parameters such as the ratios of NADH/ NAD⫹, acetyl-CoA/CoA, and ATP/ADP but also to the elaborate interactions among the component proteins of PDC such as E2, PDKs, PDPs, and E1. We now know that protein-protein interactions in PDC contribute significantly to the elaborate regulation. Moreover the structural and functional diversity of the regulatory components, PDKs and PDPs, as reviewed in this article, illustrate further the complexity necessary for tissue-specific regulation of PDC and its importance in the maintenance of glucose homeostasis in mammals. Finally, genetic defects of PDC, although rare, further illustrate the importance of PDC in both glucose and energy homeostasis not only in adult life but also from early embryonic development. Mammalian life does not exist in the complete absence of PDC! Acknowledgments—We are grateful to Dr. Richard Hanson of the Department of Biochemistry, Case Western Reserve University School of Medicine and Dr. Murray Ettinger of this department for critical reading of the manuscript and helpful discussions. REFERENCES [1] M. S. Patel, T. E. Roche, R. A. Harris, Eds. (1996) ␣-Keto Acid Dehydrogenase Complexes, Birkhauser Verlag, Basel, Switzerland. [2] M. C. Sugden, M. J. Holness (1994) Interactive regulation of the pyruvate dehydrogenase complex and the carnitine palmitoyltransferase system, FASEB J. 8, 54 – 61. [3] P. J. Randle, D. A. Priestman, S. C. Mistry, A. Halsall (1994) Glucose fatty acid interactions and the regulation of glucose disposal, J. Cell. Biochem. 55, 1–11. [4] M. S. Patel, R. A. Harris (1995) Mammalian ␣-keto acid dehydrogenase complexes: gene regulation and genetic defects, FASEB J. 9, 1164 –1172. [5] R. N. Perham (2000) Swinging arms and swinging domains in multifunctional enzymes: catalytic machines for multistep reactions, Annu. Rev. Biochem. 69, 961–1004. [6] T. E. Roche, J. C Baker, X. Yan, Y. Hiromasa, X. Gong, T. Peng, J. Dong, A. Turkan, S. A. Kasten (2001) Distinct regulatory properties of pyruvate dehydrogenase kinase and phosphatase isoforms, Prog. Nucleic Acid Res. Mol. Biol. 70, 33–75. [7] R. A. Harris, B. Huang, P. Wu (2001) Control of pyruvate dehydrogenase kinase gene expression, Adv. Enzyme Regul. 41, 269 –288. [8] L. J. Reed (2001) A trail of research from lipoic acid to ␣-keto acid dehydrogenase complexes, J. Biol. Chem. 276, 38329 –38336. [9] A. F. Hengeveld, A. deKok (2002) Structural basis of the dysfunctioning of human 2-oxo acid dehydrogenase complexes, Curr. Med. Chem. 9, 499 –520. [10] M. S. Patel, L. G. Korotchkina (2002) in Wiley Encyclopedia of Molecular Medicine, John Wiley and Sons, Inc., New York, Vol. 5, pp. 2699 –2703. [11] R. A. Harris, M. M. Bowker-Kinley, B. Huang, P. Wu (2002) Regulation of the activity of the pyruvate dehydrogenase complex, Adv. Enzyme Regul. 42, 249 –259. [12] M. S. Patel, T. E. Roche (1990) Molecular biology and biochemistry of pyruvate dehydrogenase complexes, FASEB J. 4, 3224 –3233. [13] J. K. Stoops, R. H. Cheng, M. A. Yazdi, C. Y. Maeng, J. P. Schroeter, U. Klueppelberg, S. J. Kolodziej, T. S. Baker, L. J. Reed (1997) On the unique structural organization of the Saccharomyces cerevisiae pyruvate dehydrogenase complex, J. Biol. Chem. 272, 5757–5764. [14] M. M. Klingbeil, D. J. Walker, R. Arnette, E. Sidawy, K. Hayton, P. R. Komuniecki, R. Komuniecki (1996) Identification of a novel dihydrolipoyl dehydrogenase-binding protein in the pyruvate dehydrogenase complex of the anaerobic parasitic nematode, Ascaris suum, J. Biol. Chem. 271, 5451–5457. [15] P. Arjunan, N. Nemeria, A. Brunskill, K. Chandrasekhar, M. Sax, Y. Yan, F. Jordan, J. R. Guest, W. Furey (2002) Structure of the pyruvate dehydrogenase multienzyme complex E1 component from Escherichia coli at 1.85 Å resolution, Biochemistry 41, 5213–5221. [16] S. S. Mande, S. Sarfaty, M. D. Allen, R. N. Perham, W. G. J. Hol (1996) Protein-protein interactions in the pyruvate dehydrogenase multienzyme complex: dihydrolipoamide dehydrogenase complexed with the binding domain of dihydrolipoamide acetyltransferase, Structure 4, 277–286. [17] Z. H. Zhou, W. Liao, R. H. Cheng, J. E. Lawson, D. B. McCarthy, L. J. Reed, J. K. Stoops (2001) Direct evidence for the size and conformational variability of the pyruvate dehydrogenase complex revealed by three-dimensional electron microscopy. The “breathing” core and its functional relationship to protein dynamics, J. Biol. Chem. 276, 21704 –21713. [18] Z. H. Zhou, D. B. McCarthy, C. M. O’Connor, L. J. Reed, J. K. Stoops (2001) The remarkable structural and functional organization of the eukaryotic pyruvate dehydrogenase complexes, Proc. Natl. Acad. Sci. U. S. A. 98, 14802–14807. [19] A. Aevarsson, K. Seger, S. Turley, J. R. Sokatch, W. G. Hol (1999) Crystal structure of 2-oxoisovalerate and dehydrogenase and the architecture of 2-oxo acid dehydrogenase multienzyme complexes, Nat. Struct. Biol. 6, 785–792. [20] A. Aevarsson, J. L. Chuang, R. M. Wynn, S. Turley, D. T. Chuang, W. G. J. Hol (2000) Crystal structure of human branched-chain ␣-ketoacid dehydrogenase and the molecular basis of multienzyme complex deficiency in maple syrup urine disease, Structure 8, 277–291. [21] A. Tripatara, L. G. Korotchkina, M. S. Patel (1999) Characterization of point mutations in patients with pyruvate dehydrogenase deficiency: role of methionine-181, proline-188, and arginine-349 in the ␣ subunit, Arch. Biochem. Biophys. 367, 39 –50. [22] S. J. Jacobia, L. G. Korotchkina, M. S. Patel (2001) Differential effects of two mutations at arginine-234 in the ␣ subunit of human pyruvate dehydrogenase, Arch. Biochem. Biophys. 395, 121–128. [23] N. Nemeria, A. Volkov, A. Brown, J. Yi, L. Zipper, J. R. Guest, F. Jordan (1998) Systematic study of the six cysteines of the E1 subunit of the pyruvate dehydrogenase multienzyme complex from Escherichia coli: none is essential for activity, Biochemistry 37, 911–922. [24] H.-H. M. Dahl, J. Fitzgerald, R. Iannelo, M. S. Patel, T. E. Roche, R. A. Harris, Eds. (1996) ␣-Keto Acid Dehydrogenase Complexes, Birkhauser Verlag, Basel, Switzerland, pp. 213–226. [25] Y. J. Huang, D. Walker, W. Chen, M. Klingbeil, R. Komuniecki (1998) Expression of pyruvate dehydrogenase isoforms during the aerobic/ anaerobic transition in the development of the parasitic nematode Ascaris suum: altered stoichiometry of phosphorylation/inactivation, Arch. Biochem. Biophys. 352, 263–270. [26] L. G. Korotchkina, M. S. Patel (2001) Site specificity of four pyruvate dehydrogenase kinase isoenzymes toward the three phosphorylation sites of human pyruvate dehydrogenase, J. Biol. Chem. 276, 37223–37229. [27] M. S. Patel, L. G. Korotchkina (2001) Regulation of mammalian pyruvate dehydrogenase complex by phosphorylation: complexity of multiple phosphorylation sites and kinases, Exp. Mol. Med. 33, 191–197. [28] R. A. Harris, K. M. Popov, Y. Zhao, N. Y. Kedishvili, Y. Shimomura, D. W. Crabb (1995) A new family of protein kinases—the mitochondrial protein kinases, Adv. Enzyme Regul. 35, 147–162. [29] C. N. Steussy, K. M. Popov, M. M. Bowker-Kinley, R. B. Sloan, R. A. Harris, J. A. Hamilton (2001) Structure of pyruvate dehydrogenase kinase. Novel folding pattern for a serine protein kinase, J. Biol. Chem. 276, 37443–37450. [30] L. G. Korotchkina, M. S. Patel (2001) Probing the mechanism of inactivation of human pyruvate dehydrogenase by phosphorylation of 15 three sites, J. Biol. Chem. 276, 5731–5738. [31] M. Caruso, M. A. Maitan, G. Bifulco, C. Miele, G. Vigliotta, F. Oriente, P. Formisano, F. Beguinot (2001) Activation and mitochondrial translocation of protein kinase C␦ are necessary for insulin stimulation of pyruvate dehydrogenase complex activity in muscle and liver cells, J. Biol. Chem. 276, 45088 – 45097. [32] M. H. Luethy, J. A. Miernyk, N. R. David, D. D. Randall, in M. S. Patel, T. E. Roche, R. A. Harris, Eds. (1996) ␣-Keto Acid Dehydrogenase Complexes, Birkhauser Verlag, Basel, Switzerland, pp. 71–92. [33] M. S. Patel, S. Naik, M. Johnson, R. Dey, in M. S. Patel, T. E. Roche, R. A. Harris, Eds. (1996) ␣-Keto Acid Dehydrogenase Complexes, Birkhauser Verlag, Basel, Switzerland, pp. 197–211. [34] L. A. Da Silva, O. L. De Marcucci, Z. R. Kuhnle (1993) Dietary polyunsaturated fats suppress the high-sucrose-induced increase of rat liver pyruvate dehydrogenase levels, Biochim. Biophys. Acta 1169, 126 –134. [35] J. Tan, H. S. Yang, M. S. Patel (1998) Regulation of mammalian pyruvate dehydrogenase ␣ subunit gene expression by glucose in HepG2 cells, Biochem. J. 336, 49 –56. [36] M. C. Sugden, H. S. Lall, R. A. Harris, M. J. Holness (2000) Selective modification of the pyruvate dehydrogenase kinase isoform profile in skeletal muscle in hyperthyroidism: implications for the regulatory impact of glucose on fatty acid oxidation, J. Endocrinol. 167, 339 –345. [37] M. J. Holness, N. D. Smith, K. Bulmer, T. Hopkins, G. F. Gibbons, M. C. Sugden (2002) Evaluation of the role of peroxisome-proliferatoractivated receptor ␣ in the regulation of cardiac pyruvate dehydrogenase kinase 4 protein expression in response to starvation, high-fat feeding and hyperthyroidism, Biochem. J. 364, 687– 694. [38] B. Huang, K. M. Popov, R. A. Harris (2002) Expression of pyruvate [39] [40] [41] [42] [43] [44] [45] dehydrogenase phosphatase 2 is decreased in diabetes and starvation, Diabetes 51, Suppl. 2, A136 (Abstr. 1288P). M. Chang, S. Naik, G. L. Johanning, L. Ho, M. S. Patel (1993) Multiple protein-binding domains and functional cis-elements in the 5⬘-flanking region of the human pyruvate dehydrogenase ␣-subunit gene, Biochemistry 32, 4263– 4269. R. Dey, S. Naik, M. S. Patel (1996) Tissue-specific expression of the human dehydrogenase a (Pdha-1)/chloramphenicol acetyltransferase fusion gene in transgenic mice, Biochim. Biophys. Acta 1305, 189 –195. R. C. Iannello, I. Kola, H. H. Dahl (1993) Temporal and tissue-specific interactions involving novel transcription factors and the proximal promoter of the mouse Pdha-2 gene, J. Biol. Chem. 268, 22581–22590. U. Datta, I. D. Wexler, D. S. Kerr, I. Raz, M. S. Patel (1999) Characterization of the regulatory region of the human testis-specific form of the pyruvate dehydrogenase a-subunit (PDHA-2) gene, Biochim. Biophys. Acta 1447, 236 –243. D. S. Kerr, I. D. Wexler, A. Tripatara, M. S., Patel, in M. S. Patel, T. E. Roche, R. A. Harris, Eds. (1996) ␣-Keto Acid Dehydrogenase Complexes, Birkhauser, Basel, Switzerland, pp. 249 –269. W. Lissens, L. De Meirleir, S. Seneca, I. Liebaers, G. K. Brown, R. M. Brown, M. Ito, E. Naito, Y. Kuroda, D. S. Kerr, I. D. Wexler, M. S. Patel, B. H. Robinson, A. Seyda (2000) Mutations in the X-linked pyruvate dehydrogenase (E1) ␣ subunit gene (PDHA1) in patients with a pyruvate dehydrogenase complex deficiency, Hum. Mutat. 15, 209 –219. M. T. Johnson, S. Mahmood, S. L. Hyatt, H. S. Yang, P. D. Soloway, R. W. Hanson, M. S. Patel (2001) Inactivation of the murine pyruvate dehydrogenase (Pdha1) gene and its effect on early embryonic development, Mol. Genet. Metab. 74, 293–302.