e1 enzyme of the pyruvate dehydrogenase complex in ...e1 enzyme of the pyruvate dehydrogenase...

JOURNAL OF BACTERIOLOGY, Sept. 2005, p. 6005–6018 Vol. 187, No. 170021-9193/05/$08.00�0 doi:10.1128/JB.187.17.6005–6018.2005Copyright © 2005, American Society for Microbiology. All Rights Reserved.

E1 Enzyme of the Pyruvate Dehydrogenase Complex inCorynebacterium glutamicum: Molecular Analysis of the

Gene and Phylogenetic AspectsMark E. Schreiner, Diana Fiur, Jirı Holatko,† Miroslav Patek,†

and Bernhard J. Eikmanns*Department of Microbiology and Biotechnology, University of Ulm, 89069 Ulm, Germany

Received 22 March 2005/Accepted 10 June 2005

The E1p enzyme is an essential part of the pyruvate dehydrogenase complex (PDHC) and catalyzes theoxidative decarboxylation of pyruvate with concomitant acetylation of the E2p enzyme within the complex. Weanalyzed the Corynebacterium glutamicum aceE gene, encoding the E1p enzyme, and constructed and charac-terized an E1p-deficient mutant. Sequence analysis of the C. glutamicum aceE gene and adjacent regionsrevealed that aceE is not flanked by genes encoding other enzymes of the PDHC. Transcriptional analysisrevealed that aceE from C. glutamicum is monocistronic and that its transcription is initiated 121 nucleotidesupstream of the translational start site. Inactivation of the chromosomal aceE gene led to the inability to growon glucose and to the absence of PDHC and E1p activities, indicating that only a single E1p enzyme is presentin C. glutamicum and that the PDHC is essential for the growth of this organism on carbohydrate substrates.Surprisingly, the E1p enzyme of C. glutamicum showed up to 51% identity to homodimeric E1p proteins fromgram-negative bacteria but no similarity to E1 �- or �-subunits of heterotetrameric E1p enzymes which aregenerally assumed to be typical for gram-positives. To investigate the distribution of E1p enzymes in bacteria,we compiled and analyzed the phylogeny of 46 homodimeric E1p proteins and of 58 �-subunits of heterotet-rameric E1p proteins deposited in public databases. The results revealed that the distribution of homodimericand heterotetrameric E1p subunits in bacteria is not in accordance with the rRNA-based phylogeny of bacteriaand is more heterogeneous than previously assumed.

The pyruvate dehydrogenase complex (PDHC) repre-sents a member of a multienzyme complex family that alsocomprises the 2-oxoglutarate dehydrogenase complex(OGDHC) and the branched-chain 2-oxoacid dehydroge-nase complex (BCOADHC). These enzymes catalyze theoxidative decarboxylation of pyruvate, 2-oxoglutarate, andthe 2-oxo acids of the branched-chain amino acids L-leucine,L-valine, and L-isoleucine, respectively. In general, the mul-tienzyme complexes are composed of multiple copies ofthree different enzymes, a thiamine pyrophosphate (TPP)containing 2-oxoacid decarboxylase (E1), a lipoic acid-con-taining dihydrolipoamide acyltransferase (E2), and the fla-vin-containing lipoamide dehydrogenase (LPD). The E1 en-zyme catalyzes the irreversible, TPP-dependent oxidativedecarboxylation of the 2-oxoacid, followed by the acylationof the lipoyl prosthetic group covalently attached to the E2chain. The E2 component catalyzes the transfer of the acylgroup from the lipoyl group to coenzyme A (CoA). Theresulting dihydrolipoyl group is reoxidized by LPD, gener-ating NADH and H� from NAD� (for a recent review, seereference 11). The E1 and E2 enzymes are specific for eachof the three multienzyme complexes and therefore specified

as E1p and E2p in the PDHC, E1o and E2o in the OGDHC,and E1b, and E2b in the BCOADHC. In contrast, the LPDcomponent is common in the three multienzyme complexesin most organisms (11, 42). Depending on the organism andthe type of 2-oxoacid dehydrogenase complex, the E1 en-zyme exists either as a homodimer (�2) or as a heterotet-ramer (�2�2), the subunits of both types showing only veryweak similarity to each other and not being related (11). Inall known OGDHCs and in the PDHCs of all gram-negativebacteria investigated, the E1 enzyme represents a ho-modimeric enzyme, whereas in all known BCOADHCs andin the PDHCs of gram-positive bacteria and eukaryotes, E1represents a heterotetramer (17, 28, 37, 42). The knownexceptions from this rule are Zymomonas mobilis and Thio-bacillus ferrooxidans, both of which are gram-negatives andhave been shown to possess heterotetrameric E1p enzymes(37).

The genes encoding the E1p enzymes from a number ofdifferent bacteria have been cloned and (functionally) charac-terized (reviewed in reference 37). They are designated eitheraceE or pdhA in the case of the homodimeric E1p enzyme oras pdhA(�) and pdhA(�) in the case of the heterotetramericE1p �- and �-subunits, respectively. In most organisms, thegene(s) encoding the E1p components is clustered with thegene encoding the E2 subunit of the PDHC (i.e., depending onthe organism, it is designated the aceF, pdhB, or pdhC gene[37]). The LPD gene(s) in most cases is located either togetherwith the E1p or E2p gene or in the cluster of the odh genes,which encode the enzyme components of the OGDHC (24, 37,53, 61, 72, 73).

* Corresponding author. Mailing address: Department of Microbi-ology and Biotechnology, University of Ulm, 89069 Ulm, Germany.Phone: 49 (0) 731 50 22707. Fax: 49 (0) 731 50 22719. E-mail:[email protected].

† Present address: Institute of Microbiology, Academy of Sciencesof the Czech Republic, Vıdenska 1083, CZ-14220 Prague 4, CzechRepublic.

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Corynebacterium glutamicum is an aerobic, gram-positive or-ganism that grows on a variety of sugars and organic acids andis widely used in the industrial production of amino acids,particularly L-glutamate and L-lysine (35). Due to its impor-tance for the carbon flux distribution within metabolism andfor the precursor supply for amino acid synthesis, the phos-phoenolpyruvate-pyruvate node of C. glutamicum has beenintensively studied and much attention has been focused onsome of the enzymes involved (reviewed in reference 49).However, despite its crucial role, the PDHC of this organismhas only scarcely been investigated at the molecular and struc-tural levels. The activity of the PDHC has been detected invarious strains of C. glutamicum (8, 9, 56, 59). The activitydepends on the substrate pyruvate and the cofactors TPP andMg2� and is higher in the presence of either cysteine or di-thiothreitol (59). According to enzyme measurements with cellextracts, the C. glutamicum PDHC is not subject to any signif-icant regulation modulating its activity (8, 9, 64). This seemssurprising in view of the complex regulation of PDHCs in otherbacteria as well as in eukaryotes (5, 18, 20, 23, 58). Whereas theC. glutamicum E1p and E2p proteins and their genes have notbeen investigated so far, Schwinde et al. (56) cloned and ana-lyzed a functional lpd gene from C. glutamicum and purifiedand biochemically characterized the corresponding LDP pro-tein. The lpd gene is monocistronic and not clustered with thegenes for other enzymes of the PDHC or of the OGDHC.However, as lpd-deficient mutants of C. glutamicum have not

yet been generated and analyzed, it remains unclear whetherthe characterized LPD protein functions as the third subunit ofthe PDHC and/or of another 2-oxoacid dehydrogenase com-plex in C. glutamicum.

The genome sequence of C. glutamicum has recently beendetermined and annotated (GenBank accession numbersNC_003450 and BX927147) (26, 29, 62), and an open readingframe (cg2466) coding for a protein with significant similarityto the Escherichia coli E1p enzyme has been detected andaccordingly designated the aceE gene (29). In the present studywe describe the genetic and functional characterization of theC. glutamicum E1 enzyme of the PDHC. It represents the firstexample for a homodimeric E1p protein in a gram-positivebacterium. Furthermore, we performed sequence analyses ofall E1 amino acid sequences available in public databases andinvestigated the distribution as well as the phylogeny of ho-modimeric and heterotetrameric E1p proteins in prokaryotes.

MATERIALS AND METHODS

Bacteria, plasmids, oligonucleotides, and culture conditions. All bacterialstrains and plasmids and their relevant characteristics and sources are given inTable 1. The oligonucleotides used and their sequences are also listed in Table1. The minimal medium used for C. glutamicum has been described previously(14) and contained 1% (wt/vol) acetate, lactate, pyruvate, or 2% (wt/vol) glucoseas the carbon and energy source. Tryptone-yeast extract (TY) medium (48) wasused as the complex medium for C. glutamicum and E. coli. When appropriate,kanamycin (50 �g ml�1) was added to the medium. If not stated otherwise, C.glutamicum was grown aerobically at 30°C and E. coli was grown aerobically at

TABLE 1. Strains, plasmids and oligonucleotides used in this study

Strain, plasmid, oroligonucleotide Relevant characteristic(s) or sequence Source/reference or purpose

StrainsE. coli DH5� supE44 hsdR17 recA1 endA1 gyrA96 thi-1 relA1 21

C. glutamicumWT WT strain ATCC 13032 American Type Culture CollectionWT �aceE mutant WT strain with deletion of aceE, encoding the E1 enzyme of the PDHC This work

PlasmidspK19mobsacB Kmr, mobilizable (oriT), oriV 50pK19mutaceE pK19mobsacB carrying a 1,155-bp insert with a truncated aceE gene,

shortened by 2,077 bpThis work

pET2 Promoter probe vector carrying the promoterless cat gene, Kmr 66pET-PaceE pET2 containing a 409-bp insert of the aceE promoter region This work

OligonucleotidesdelaceE1 5�-ACGCGTCGACCACCAAAAGGACATCAGACC-3� Primer for deletion in aceE, SalI

site (underlined)delaceE3 5�-GGATAGGTGATTGGAAGTTGGGCAAACGAAGCATGAGGTAACG-3� Primer for deletion in aceE,

crossover overlap (italicized)delaceE4.1 5�-CCAACTTCCAATCACCTATCCGGGCATCTACCTCTACTC-3� Primer for deletion in aceE,

crossover overlap (italicized)delaceE5.3 5�-CGCGGATCCGCGGGATTTATCTGTCCC-3� Primer for deletion in aceE,

BamHI site (underlined)aceEprom1 5�-ACGCGTCGACCACCAAAAGGACATCAGACC-3� Primer for amplification of the

aceE promotor, SalI site(underlined)

aceEprom2 5�-CGCGGATCCACACCTCCTGTTGGAATG-3� Primer for amplification of theaceE promotor, BamHI site(underlined)

CM4 5�-GAAAATCTCGTCGAAGCTCG-3� 67CM5 5�-AAGCTCGGCGGATTTGTC-3� This work

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37°C as 60-ml cultures in 500-ml baffled Erlenmeyer flasks on a rotary shaker at120 rpm.

DNA preparation and transformation. The isolation of chromosomal DNAand plasmids from C. glutamicum was performed as described previously (15).Plasmid isolation from E. coli was carried out according to the method ofBirnboim (4). DNA transfer into C. glutamicum was performed by electropora-tion, and the recombinant strains were selected on LBBHIS agar plates contain-ing kanamycin (15 �g ml�1) (65). Electroporation of E. coli was performed withcompetent cells according to the method of Dower et al. (12).

PCR techniques. PCR experiments were performed in a Biometra personalcycler (Biotron, Gottingen, Germany). Amplification of DNA was carried outwith Vent polymerase (New England Biolabs). Buffers and deoxynucleosidetriphosphates were taken from MBI-Fermentas (St. Leon-Rot, Germany). Oli-gonucleotides (primers) were obtained from MWG-Biotech (Ebersberg, Ger-many). Cycling times and temperatures were chosen according to fragmentlength and primer constitution. PCR products were purified from agarose gelsusing the Nucleospin extract II kit (Macherey & Nagel, Duren, Germany).

DNA manipulation and Southern hybridization. Restrictions enzymes, T4DNA ligase, calf intestinal phosphatase, RNase A, proteinase K, and Taq poly-merase were purchased from MBI-Fermentas and used according to the instruc-tions of the manufacturer. After restriction digests, DNA was separated onagarose gels and purified with the Nucleospin extract II kit. DNA hybridizationexperiments were performed as previously described (15). An aceE-specific580-bp DNA fragment was amplified from chromosomal DNA of wild-type(WT) C. glutamicum by PCR with the primers delaceE1 and delaceE3 and usedas a probe. Labeling, hybridization, washing, and detection were conducted usingthe nonradioactive DNA labeling and detection kit and the instructions fromRoche Diagnostics (Penzberg, Germany).

Construction of a C. glutamicum aceE deletion mutant. Inactivation of thechromosomal aceE gene in C. glutamicum was performed as described previously(38), using crossover PCR and the suicide vector pK19mobsacB. aceE-specificDNA fragments were generated using the primer pairs delaceE1-delaceE3 anddelaceE4.1-delaceE5.3. Fragment 1 covers 393 bp upstream of aceE and 187 bpof the 5� end of aceE, and fragment 2 covers 547 bp of the 3� end of aceE and 28bp downstream of the aceE stop codon (see Fig. 2). The two fragments werepurified, mixed in equal amounts, and subjected to crossover PCR using primersdelaceE1 and delaceE5.3. The resulting fusion product (containing the aceEgene with a deletion of 2,077 bp) was digested with BamHI/SalI, ligated into theBamHI/SalI-restricted plasmid pK19mobsacB, and transformed into E. coli. Therecombinant plasmid was isolated from E. coli and electroporated into WT C.glutamicum. By application of the method described by Schafer et al. (50), theintact chromosomal aceE gene in WT C. glutamicum was replaced by the trun-cated aceE gene via homologous recombination (double crossover). The screen-ing of the aceE mutants was done on LB agar plates containing 0.5% (wt/vol)glucose and 10% (wt/vol) sucrose. The replacement at the chromosomal locuswas verified by PCR using primers delaceE1/delaceE5.3 and by Southern blotanalysis. For the latter, a labeled aceE probe was hybridized to SalI-restrictedand size-fractionated chromosomal DNA from WT C. glutamicum and the C.glutamicum �aceE mutant, resulting in one signal of about 8.3 kb with the DNAfrom the WT strain and one signal of about 6.2 kb with the DNA from the �aceEmutant.

Cloning of the aceE promoter. The aceE promoter fragment was amplifiedfrom chromosomal DNA of WT C. glutamicum by PCR with the primers aceE-prom1 and aceEprom2. The PCR product was digested with SalI and BamHI,ligated into SalI/BamHI-restricted plasmid pET2, and transformed into E. coli.The recombinant plasmid pET-PaceE was then isolated from E. coli and intro-duced into C. glutamicum by electroporation.

RNA techniques. For RNA isolation, C. glutamicum cells grown to the expo-nential phase (optical density at 600 nm of about 3) were treated with 1 volumeof ice-cold killing buffer (20 mM Tris-HCl, pH 8.0, 20 mM NaN3, MgCl2),harvested, and resuspended in 100 �l ice-cold RNase-free water. The cell sus-pension was transferred to 2-ml screw-cap vials containing 500 mg glass beads(150 to 212 �m; Sigma), 0.5 ml RNeasy lysis buffer (QIAGEN, Hilden, Ger-many), and 0.5 ml acidic phenol (pH 5.5) and then mechanically disrupted byincubation three times for 45 s at 4°C in a RiboLyser (Hybaid, Heidelberg,Germany) at setting 6.5. After disruption, glass beads and cellular debris wereremoved by centrifugation (10,000 � g, 4°C). The supernatant was extracted withhot phenol as described by Schwinde et al. (55), and then the RNA was ethanolprecipitated and dissolved in 400 �l RNase-free water. After treatment with 30units RNase-free DNase (MBI Fermentas) for 15 min at 37°C, the sample wasapplied to an RNeasy Midi spin column (QIAGEN) and purified by following theinstructions of the manufacturer. Aliquots of RNA were stored at �70°C untiluse.

For Northern (RNA) hybridization, a digoxigenin-dUTP-labeled aceE-specific580-bp DNA probe was generated as described above. For hybridization, 10 �gof total RNA from WT C. glutamicum was separated on an agarose gel contain-ing 17% (vol/vol) formaldehyde and transferred onto a nylon membrane (15).Hybridization (at 50°C, in the presence of 50% formamide, vol/vol), washing, anddetection were carried out using a nucleic acid detection kit according to theinstructions from Roche Diagnostics (Penzberg, Germany). The size marker wasthe 0.24- to 9.5-kb RNA ladder from GibcoBRL.

Primer extension reactions were carried out as described previously (30) withIRD800-labeled primers CM4 and CM5 and RNA from C. glutamicum (pET-PaceE). The IRD800-labeled primers were obtained from MWG Biotech. Prim-ers CM4 and CM5 are complementary to regions from nucleotides (nt) 45 to 64and 34 to 51, respectively, downstream of the BamHI site of the multiple cloningsite in plasmid pET2. Primer extension products were analyzed with an auto-matic sequencer (LI-COR 4000L; Licor, Inc.) using a 6% (wt/vol) polyacryl-amide gel at 1,500 V and 50°C. For the exact localization of the transcriptionalstart site, sequencing reactions using plasmid pET-PaceE and the same oligo-nucleotide used for the respective primer extension reaction were coelectro-phoresed.

Enzyme assays. To determine PDHC activity in cell extracts, C. glutamicumcells were harvested in the exponential growth phase (optical density at 600 nmof about 8), washed twice in 100 mM Tris-HCl, pH 7.2, 3 mM L-cysteine, 10 mMMgCl2, and resuspended in 0.5 ml of the same buffer. The cell suspension wastransferred to 2-ml screw-cap vials together with 250 mg glass beads (150 to 212�m; Sigma) and subjected five times for 30 s to mechanical disruption with aRiboLyser at 4°C, with intermittent cooling on ice for 2 min. After disruption, theglass beads and the cellular debris were removed by centrifugation (10,000 � g,4°C, 15 min) and the supernatant was used for the assay. The protein concen-tration was determined by a bicinchoninic acid protein assay reagent kit (Pierce,Bonn, Germany) with bovine serum albumin as the standard. PDHC enzymeactivities were determined photometrically according to the method described byGuest and Creaghan (19). One unit of activity is defined as 1 �mol NADHformed per min at 30°C.

For the determination of chloramphenicol acetyltransferase (CAT) activity,crude extracts were prepared as described above, except that cells were washedtwice in 50 mM Tris-HCl, pH 7, and resuspended in 0.5 ml of the same buffercontaining 10 mM MgCl2, 1 mM EDTA, and 30% (vol/vol) glycerol. CAT activitywas assayed photometrically at 412 nm as described by Shaw (57) in 1 ml 100 mMTris-HCl, pH 7.8, 1 mM 5,5�-dithiobis-2-nitrobenzoic acid, 0.1 mM acetyl-CoA,and 0.25 mM chloramphenicol. One unit of CAT activity is defined as 1 �molchloramphenicol acetylated per min at 37°C.

For the determination of E1p activity, crude extracts were prepared as de-scribed above, except that cells were washed twice in 50 mM Tris-HCl, pH 7.5,10% (vol/vol) glycerol, and resuspended in 0.5 ml of the same buffer. E1p activitywas assayed photometrically according to the method described by Schwartz andReed (54). One unit of activity corresponds to 2 �mol ferricyanide reduced permin at 30°C.

Computational analysis. MFold (75) was the software used for the calculationof the �Go� value (free energy under standard conditions) of the aceE terminatorstructure. Databank searches and alignments were carried out by using BLASTand CLUSTALW (2, 63). Phylogenetic trees were generated using the neighbor-joining method (46), including 100 bootstrap replicates. The accession numbersfor all homodimeric E1p and heterotetrameric E1p� and E1b� sequences usedfor the phylogenetic analysis were as follows. The homodimeric E1p sequenceswere Acinetobacter sp. strain ADP1 (YP_047975), Actinobacillus pleuropneumo-niae (ZP_00134360), Anopheles gambiae (EAL42079), Azotobacter vinelandii(ZP_00342533), Blochmannia floridanus (NP_878459), Bordetella parapertussis(NP_883760), Buchnera aphidicola (NP_660552), Burkholderia cepacia(ZP_00212749), Chromobacterium violaceum (AAQ58203), Corynebacterium glu-tamicum (CAF20589), Coxiella burnetii (NP_819497), Dechloromonas aromatica(ZP_00150166), Deinococcus radiodurans (NP_293980), Erwinia carotovora(YP_051878), Escherichia coli (NP_414656), Haemophilus influenzae(ZP_00154971), Legionella pneumophila (YP_126868), Leifsonia xyli(YP_062215), Mannheimia succiniciproducens (YP_088528), Methylobacillusflagellatus (ZP_00172315), Methylococcus capsulatus (YP_115388), Microbulbiferdegradans (ZP_00315287), Mycobacterium tuberculosis (NP_336771), Neisseriameningitidis (NP_274360), Nitrosomonas europaea (NP_840448), Nocardia far-cinica (YP_117823), Pasteurella multocida (NP_245832), Photobacterium profun-dum (YP_131304), Photorhabdus luminescens (CAE15996), Polaromonas sp.strain JS666 (ZP_00362465), Propionibacterium acnes (YP_055700), Pseudomo-nas aeruginosa (NP_253702), Psychrobacter sp. strain 273-4 (ZP_00146054), Ral-stonia metallidurans (ZP_00271472), Rhodopirellula baltica (NP_865502), Rubri-vivax gelatinosus (ZP_00245303), Salmonella enterica serovar Typhimurium

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(NP_459157), Shewanella oneidensis (NP_716061), Shigella flexneri (NP_706068),Streptomyces coelicolor (NP_626618), Thermus thermophilus (YP_005770), Tro-pheryma whipplei (NP_789444), Vibrio cholerae (AAF95557), Wautersia eutropha(AAA21598), Xanthomonas axonopodis (NP_640929), Xylella fastidiosa(NP_297959), and Yersinia pestis (NP_406881). The heterotetrameric E1p� se-quences were Acholeplasma laidlawii (P35485), Agrobacterium tumefaciens(NP_532119), Arabidopsis thaliana (AAB86803), Azorhizobium caulinodans(AAG38097), Bacillus subtilis (DEBSPA), Bartonella henselae (YP_033409), Bra-dyrhizobium japonicum (NP_771423), Brucella melitensis (NP_539771), Candidaalbicans (EAK96452), Caulobacter crescentus (NP_420534), Coxiella burnetii(NP_819670), Cyanidium caldarium (AAF12897), Cytophaga hutchinsonii(ZP_00308483), Danio rerio (NP_001002399), Enterococcus faecalis(AAO81144), Exiguobacterium sp. (ZP_00182969), Geobacter sulfurreducens(NP_953699), Gloeobacter violaceus (NP_924475), Gracilaria tenuistipitata(YP_063628), Haloferax volcanii (AAD34202), Lactobacillus plantarum(NP_785659), Lactococcus lactis (NP_266218), Leuconostoc mesenteroides(ZP_00062839), Listeria monocytogenes (NP_464577), Magnetospirillum magne-totacticum (ZP_00208699), Mesorhizobium sp. (ZP_00196269), Methylobacteriumextorquens (AAN03811), Mus musculus (NP_032837), Mycoplasma capricolum(AAC44342), Nostoc sp. (NP_486748), Novosphingobium aromaticivorans(ZP_00303573), Oceanobacillus iheyensis (NP_692333), Oenococcus oeni(ZP_00320046), Pediococcus pentosaceus (ZP_00323580), Porphyra purpurea(P51267), Prochlorococcus marinus (NP_875753), Rattus norvegicus(NP_446446), Rhodobacter sphaeroides (ZP_00007453), Rhodopseudomonaspalustris (NP_948208), Rhodospirillum rubrum (ZP_00268857), Rickettsia rickett-sii (ZP_00153395), Saccharomyces cerevisiae (NP_011105), Silicibacter sp.(ZP_00339083), Sinorhizobium meliloti (NP_385551), Staphylococcus aureus(YP_040480), Synechococcus elongatus (YP_172860), Synechocystis sp.(BAA18592), Thermosynechococcus elongatus (BAC08721), Thermus thermophi-lus (YP_144205), Trichodesmium erythraeum (ZP_00327615), and Zymomonasmobilis (AAC70361). The BCOADHC E1� sequences were B. subtilis (C69593),L. monocytogenes (NP_464897), M. musculus (NP_031559), O. iheyensis(NP_692787), Pseudomonas putida (P09060), Streptomyces avermitilis(AAA66072), and T. thermophilus (1UMD_C).

RESULTS

PDHC activity in C. glutamicum. The specific activity of thePDHC was determined in cell extracts of WT C. glutamicumgrown in TY complex medium and in minimal medium con-taining glucose or acetate as the carbon source and harvestedat the mid-exponential growth phase. The highest specific ac-tivity (0.087 U mg protein�1) was found in cells grown in TYmedium (Table 2). The activity was about two- to threefoldlower when the cells were grown in media containing glucoseor acetate. The dependency of the PDHC activity on thegrowth phase was elucidated by measuring the specific activityin cell extracts from WT C. glutamicum grown to the stationary

growth phase in TY medium and harvested in 2-hour intervals(Fig. 1). The highest specific activity was found when the cellswere harvested in the mid- to late exponential growth phase;much lower activities were found in cells of the stationaryphase. These results suggest that the PDHC in C. glutamicumis weakly regulated by the growth medium and is significantlyregulated dependent on the growth phase. In view of the cen-tral position of the PDHC in the metabolism, this regulationmight be significant for the carbon flux in C. glutamicum.

The analysis of the PDHC activity in extracts of WT C.glutamicum revealed an apparent Km value of about 1.7 mMfor pyruvate, which is about twofold higher than that (0.8 mM)reported by Shiio et al. (59). In accordance with these authors,the WT C. glutamicum enzyme required Mg2� and TPP formaximal activity.

Nucleotide sequence of the C. glutamcium aceE gene andanalysis of the deduced E1p amino acid sequence. Due tosequence similarity to the E. coli aceE gene, the cg2466 gene ofthe C. glutamicum genome has recently been annotated as anaceE gene, putatively coding for the E1 enzyme of the C.glutamicum PDHC (29). The aceE gene of C. glutamcium con-sists of 2,766 bp and is preceded by a typical ribosomal bindingsite (AGGAGG). Centered 38 bp downstream of the aceE stopcodon, a region of dyad symmetry followed by several T resi-dues similar to rho-independent transcription terminators wasfound. The mRNA hairpin loop predicted from this sequencehas a �Go� value of �24.0 kcal/mol at 25°C. This result indi-cates transcriptional termination downstream of the aceEgene. According to the C. glutamicum genome sequence (Gen-Bank accession numbers NC_003450 and BX927147 [26, 29]),three divergently orientated open reading frames, cg2467,cg2468, and cg2470, are located downstream of the aceE gene(Fig. 2). The deduced amino acid sequences of these openreading frames show significant similarities to components of

FIG. 1. Growth (black squares) and pyruvate dehydrogenase com-plex (PDHC) activity (black bars) of WT C. glutamicum grown on TYcomplex medium. The standard deviations of the enzyme determina-tions were below 10%. OD600, optical density at 600 nm.

TABLE 2. Specific activities of the PDHC in cell extracts of WTC. glutamicum and the C. glutamicum �aceE mutant grown on

different mediaa

Medium

Sp act of PDHC (U/mg protein)b

C. glutamicum C. glutamicum �aceEmutant

TY 0.087 NDTY � glucose 0.029 NDTY � acetate 0.032 0.001MM � glucose 0.036 NGMM � acetate 0.034 0.001MM � glucose � acetate 0.034 0.001

a TY medium without and with 1% glucose or 0.5% acetate; minimal medium(MM) containing 2% glucose, 1% acetate, or 1% glucose plus 1% acetate as thecarbon source. The cells were harvested in the mid-exponential growth phase.

b The values are means obtained from at least two independent cultivationsand two determinations per experiment. The standard deviations were in allcases below 10%. ND, not determined; NG, no growth.



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ABC transporters (ATP binding protein, permease, and sub-strate binding protein, respectively). Two open reading frames(cg2464 and cg2465) were identified in the region up to 1,000bp upstream of the aceE gene, both annotated as coding forhypothetical (unknown) proteins. According to the nucleotidesequence, the C. glutamicum aceE gene encodes a polypeptideof 922 amino acids with a predicted molecular mass of 102.8kDa. Alignment studies revealed the 49% to 51% identity ofthe C. glutamicum protein to the functionally well-character-ized homodimeric E1 enzymes of PDHCs from the gram-neg-atives E. coli, Wautersia (Ralstonia) eutropha, Azotobacter vine-landii, Pseudomonas aeruginosa, and Neisseria meningitidis(Fig. 3). The alignment of the C. glutamicum enzyme with thehomodimeric E1p proteins revealed highly conserved residuesand sequence motifs, especially in those regions suggested tobe essential for cofactor binding and catalytic activity (3).These motifs include the TPP binding motif GDG. . .X26. . .NCN (residues 253 to 284 in the C. glutamicum se-quence) located in the N-terminal part of all homodimeric E1penzymes, binding sites for divalent cations (Asp254, Asn284,and Gln286), and several residues proposed to be involved incofactor binding and catalysis (His128, His164, His668,Glu550, Tyr201, and Asp549) (Fig. 3). Surprisingly, the data-bank searches of the C. glutamicum genome revealed no pu-tative proteins showing similarity to the E1� or E1� subunits ofheterotetrameric E1p enzymes, generally assumed to be typicalfor gram-positive bacteria (11, 17, 37). In summary, sequenceanalysis of the cg2466 gene suggests the presence of a ho-modimeric E1p enzyme in the PDHC of gram-positive C. glu-tamicum.

Inactivation of the chromosomal aceE gene. To studywhether C. glutamicum requires the aceE gene for growth andfor PDHC activity, the chromosomal aceE locus was partiallydeleted and the resulting strain, the C. glutamicum �aceEmutant, was tested for growth in different media and forPDHC activity. The growth of the C. glutamicum �aceE mu-tant was negligible in TY medium, but the strain grew as wellas WT C. glutamicum on TY medium supplemented with ac-etate. Furthermore, the C. glutamicum �aceE mutant was un-able to grow in minimal medium with glucose, pyruvate, orlactate as the sole carbon source but grew almost identically toWT C. glutamicum in minimal medium containing glucose plusacetate as carbon sources (Fig. 4). In accordance with thesefindings, the growth of the C. glutamicum �aceE mutant wasindistinguishable from that of the WT strain in minimal me-

dium containing acetate as the sole carbon source (Fig. 4).These results indicate that the aceE gene is essential for thegrowth of C. glutamicum in minimal medium containing glu-cose or other substrates entering the central metabolism asintermediates of glycolysis but not for growth in minimal me-dium with acetate as the carbon and energy source.

To investigate whether the aceE gene in fact encodes the E1enzyme of the PDHC in C. glutamicum, both the specificPDHC and E1p activities were determined in cell extracts ofWT C. glutamicum and the C. glutamicum �aceE mutantgrown in complex and minimal media containing acetate. Asshown in Table 2, the C. glutamicum �aceE mutant was devoidof any detectable PDHC activity (0.01 U mg protein�1),whereas the WT strain showed 0.032 U mg protein�1 and 0.034U mg protein�1, respectively. Furthermore, no E1p activitiy(0.01 U mg protein�1) was detected in extracts of C. glutami-cum �aceE mutant cells grown in TY medium supplementedwith acetate (0.5%, wt/vol), whereas an activity of 0.017 U mgprotein�1 was measured in extracts of WT C. glutamicum.These findings indicate (i) that the aceE gene in fact encodesthe E1 enzyme of the PDHC in C. glutamicum and (ii) thatthere are no isoenzymes for the E1p protein in C. glutamicum.Furthermore, our findings represent the first example of ahomodimeric E1 enzyme of a PDHC in a gram-positive bac-terium.

Transcriptional analysis of the aceE gene. Northern (RNA)hybridization experiments were performed in order to analyzethe size of the aceE transcript. For this purpose, total RNAfrom WT C. glutamicum was isolated, size-fractionated, trans-ferred onto a nylon membrane, and hybridized to an aceE-specific digoxigenin-UTP-labeled DNA probe. The hybridiza-tion revealed a signal at about 2.9 kb (data not shown), whichcorresponds well to the size of the aceE gene (2.8 kb). Thisresult indicates that the C. glutamicum aceE transcript is mono-cistronic.

To confirm the presence of a promoter and to investigatetranscriptional regulation of the aceE gene, a transcriptionalfusion between the putative aceE promoter region and thepromoterless chloramphenicol acetyltransferase (CAT) genewas constructed in the promoter probe vector pET2. The re-sulting plasmid, pET2-PaceE, was transformed into WT C.glutamicum and CAT activity was determined in the plasmid-carrying strain during growth in TY medium and in minimalmedium containing glucose or acetate as the carbon source.Whereas C. glutamicum carrying the host plasmid pET2

FIG. 2. Genomic locus of the aceE gene in C. glutamicum. The arrows represent the coding regions of aceE and adjacent genes. Thetranscriptional start site and the terminator structure are indicated as Ts and T, respectively. The double-headed arrows indicate the fragmentsused for Southern and Northern blot hybridizations and for the construction of the aceE deletion mutant.



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showed no CAT activity (0.01 U mg protein�1) on eithermedium, the strain carrying pET2-PaceE showed 0.92 U mgprotein�1 during growth in TY medium and approximately1.5-fold-lower CAT activity during growth in minimal mediumcontaining glucose or acetate (0.61 and 0.58 U mg protein�1,respectively). These results confirm the presence of a promoterimmediately upstream of the aceE gene and indicate weaktranscriptional control of the aceE gene by the carbon sourcein the growth medium.

To identify the transcriptional initiation site of the C. glu-tamcium aceE gene and to localize the aceE promoter, primerextension experiments were performed. Using oligonucleotideCM4 and 50 �g total RNA isolated from C. glutamicum pET2-PaceE, a major signal corresponding to an A residue 121 nu-cleotides upstream of the aceE translational start was obtained(Fig. 5). This result was confirmed in an independent experi-ment with primer CM5 (data not shown). At an appropriatedistance (8 bp) upstream of the transcriptional initiation site ofthe aceE gene, a sequence, TATCCT, with significant similarityto the consensus �10 region (TAT/CAAT) of C. glutamicumpromoters (40, 41) is present. No apparent �35 region (TTGCCA) can be recognized, which is a common feature of C.

FIG. 4. Growth of WT C. glutamicum (filled squares) and the C.glutamicum �aceE mutant (open triangles) on minimal medium con-taining 2% glucose (A), 1% glucose plus 1% acetate (B), and 1%acetate (C). OD600, optical density at 600 nm.

FIG. 5. Primer extension analysis of the transcriptional start site infront of the aceE gene. The primer extension product is shown in lane5. Lanes A, C, G, and T represent the products of sequencing reactionswith the same primer used for the primer extension reaction. Therelevant DNA sequence (coding strand) is shown on the right, and thetranscriptional start site is indicated by an asterisk.

FIG. 3. Amino acid sequence alignment of the E1p enzymes of C. glutamicum (accession number CAF20589), E. coli (NP_414656), Wautersiaeutropha (AAA21598), Azotobacter vinelandii (ZP_00342533), Pseudomonas aeruginosa (NP_253702), and Neisseria meningitidis (NP_274360).Amino acids identical in all sequences are shaded in black, and amino acids identical in at least five sequences are shaded in gray. The TPPsignature motif is underlined.



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glutamicum promoters (40, 41). In addition to the main signalat position �121, with both primer CM4 and primer CM5 weobtained a less prominent primer extension signal which cor-responded to an A residue 54 nucleotides upstream of the aceEtranslational start site. However, analysis of the DNA se-quence upstream of this site revealed no motifs which aresimilar to the consensus �10 and �35 regions of C. glutami-cum vegetative promoters.

Sequence comparisons and phylogenetic analyses. Data-bank searches with the amino acid sequence of homodimericE1p of C. glutamicum revealed, aside from similarities to thefunctionally proven E1p proteins of different gram-negatives(Fig. 3), significant levels of identity (�40%) to putative orhypothetical E1p proteins from other bacteria, e.g., from“high-GC-content gram-positives,” deinococci, planctomyce-tes, several proteobacteria, and, surprisingly, the eukaryoticAnopheles gambiae (Table 3). To elucidate the phylogeny ofthe homodimeric E1p protein family, a phylogenetic tree basedon comparison of the deduced amino acid sequences of 46 E1pgenes currently available in public databases was constructed.As shown in Fig. 6, the phylogenetic tree of homodimeric E1pproteins contains four major clusters. Clusters I and IV pre-dominantly entail sequences of the -proteobacteria, whereascluster II comprises all sequences of the high-GC-contentgram-positives, deinococci, and planctomycetes and several se-quences of -proteobacteria and cluster III contains mainlyproteins of the �-proteobacteria.

To elucidate also the distribution of heterotetrameric E1pproteins in bacteria, databank searches were carried out usingthe functionally proven E1p �-subunits of Z. mobilis (37) andBacillus subtilis (24) (Table 4). The E1p �-subunits of Z. mo-bilis showed highest similarity (�40% identity) to putativeproteins of many �-proteobacteria, sphingobacteria, and sev-eral eukaryotes, e.g., Mus musculus, Rattus norvegicus, andSaccharomyces cerevisiae. In contrast, the B. subtilis proteinshowed only relatively weak similarity to the Z. mobilis en-zyme, but high levels of identity (�40%) to the described E1p�-subunits from Acholeplasma laidlawii (68) and to putativeE1p �-subunits from several low-GC-content gram-positives,-proteobacteria, deinococci, and the archaeon Haloferax vol-

canii. Surprisingly, the B. subtilis E1p �-subunit showed onlyweak similarity to the functionally proven protein of the low-GC-content gram-positive bacterium Mycoplasma capricolum(74).

A phylogenetic tree of the heterotetrameric E1p proteinfamily was constructed using the deduced amino acid se-quences of 58 E1p �-subunits deposited in public databases. Asdepicted in Fig. 7, this tree contains four major clusters broadlycorresponding to the low-GC-content gram-positive bacteria(cluster I), cyanobacteria, and chloroplasts (cluster II), themitochondria (cluster III), and the �-proteobacteria (clusterIV).

Surprisingly, databank searches and sequence comparisonsrevealed that the �-subunits of the known heterotetramericE1p proteins of Z. mobilis, B. subtilis, M. capricolum, A. laid-lawii, and Synechocystis sp. show significant similarity (28 to42% identity) to the known heterotetrameric E1 �-subunits ofthe BCOADHC from B. subtilis (69), S. avermitilis (60), P.putida (6), T. thermophilus (36), and M. musculus (10). Allthese BCOADHC E1 �-proteins cluster with the PDHC E1�-proteins of the low-GC-content gram-positive bacteria (Fig.7, cluster I).

In summary, the sequence comparisons and phylogeneticstudies suggest that (i) homodimeric E1p proteins are typicalfor the PDHCs of �- and -proteobacteria, the deinococci, andthe high-GC-content gram-positive bacteria and (ii) heterotet-rameric E1p proteins are typical for the PDHCs of the �-pro-teobacteria, the low-GC-content gram-positive bacteria, andthe cyanobacteria.

DISCUSSION

It has generally been assumed that aerobic gram-negativebacteria possess homodimeric and that gram-positive bacteriapossess heterotetrameric E1p enzymes in their PDHCs. Thepresent study describes for the first time the genetic and func-tional characterization of an E1p enzyme of the homodimerictype from a gram-positive bacterium, i.e., from C. glutamicum.

DNA sequence analysis of the C. glutamicum E1p gene(aceE) and adjacent open reading frames and comparisonswith the respective gene loci in other bacteria highlighted somedifferences between the chromosomal organization of the C.glutamicum aceE locus and that of functionally proven aceE(pdhA) genes in other microorganisms. Except in Z. mobilis,the genes for the (homodimeric and heterotetrameric) E1pand the E2p enzymes of the PDHCs are clustered in the ge-nomes of all bacteria studied so far (37). Additionally, a geneencoding an LPD can be found downstream of the E1p andE2p genes in some of these bacteria (37). A functional lpd gene(cg0441) putatively coding for the LPD enzyme of the PDHChas been identified in the C. glutamicum genome; however, itis not located in the region of the aceE locus and it is notclustered with genes for other subunits of the PDHC or theOGDHC (13, 56). Other genes (i.e., cg2194, cg0790, andcg3339) coding for proteins with some similarity (24 to 29%identity) to LPD proteins from other bacteria are also notclustered with the aceE gene or with a gene possibly coding foran E2 protein. A single gene (cg2421 or NCgl2126) for aprotein with homology to the E2 subunits of PDHCs orOGDHCs can be found in the C. glutamicum genome (29);

TABLE 3. Protein sequence identities among representativehomodimeric E1p enzymes

Organisma

% Identity

Escherichia coli Corynebacteriumglutamicum

Corynebacterium glutamicum 49Streptomyces coelicolor 47 61Mycobacterium tuberculosis 49 66Propionibacterium acnes 50 63Deinococcus radiodurans 48 47Thermus thermophilus 52 52Rhodopirellula baltica 55 51Escherichia coli 49Vibrio cholerae 72 51Burkholderia cepacia 59 50Coxiella burnetii 51 43Chromobacterium violaceum 62 52Anopheles gambiae 75 52

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however, so far this gene has not been studied and it remainsto be proven whether it in fact represents a functional E2 gene.Thus, although a functional E2p gene (aceF or pdhB) so far hasnot yet been identified in C. glutamicum, it is obvious fromsequence analyses up- and downstream of the aceE gene thatin this organism the genes encoding the E1 and E2 enzymes ofthe PDHC are also not clustered. In E. coli, the structuralgenes for the PDHC are transcribed in a single operon, to-gether with an upstream gene encoding the regulatory protein

PdhR (44). This regulator negatively controls the expression ofthe whole pdh operon, probably with pyruvate serving as aninducing effector (44). The C. glutamicum aceE gene is ex-pressed as an independent single cistron, and from genomeand sequence analyses, there is no indication for a PdhR ho-mologue. All these findings indicate that the chromosomalaceE locus in C. glutamicum does not resemble any of the aceEor pdhA loci analyzed so far from other organisms and that thetranscriptional organization in this organism is different.

FIG. 6. Phylogenetic tree of the homodimeric E1 protein family. Numbers at the nodes give the bootstrap values. Organisms are listed inMaterials and Methods.



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The enzymatic characterization of the defined C. glutami-cum �aceE mutant unequivocally demonstrated that the aceEgene in fact codes for the E1 subunit of the PDHC. Further-more, the analysis revealed that the PDHC is essential for thegrowth of C. glutamicum on minimal medium containing glu-cose, pyruvate, or lactate as the carbon and energy source.These findings indicate that under the conditions tested, thePDHC is the main enzyme responsible for providing acetyl-CoA during the growth of C. glutamicum on substrates orintermediates that do not enter the central metabolism viaacetyl-CoA. In contrast, the PDHC is dispensable for thegrowth of C. glutamicum on minimal medium containing ace-tate as the sole or as an additional carbon and energy source.This substrate is activated in C. glutamicum to acetyl-CoA byacetate kinase and phosphotransacetylase (45, 71) and then,independently of the PDHC, channeled into the tricarboxylicacid cycle. In accordance with our results, the known PDHC-negative mutants of E. coli, Pseudomonas aeruginosa, and Sal-monella enterica serovar Typhimurium also show an acetate-auxotrophic phenotype (1, 25, 27, 33). Very recently, weidentified, purified, and characterized a pyruvate:quinone ox-idoreductase (PQO) in C. glutamicum (52). This enzyme is ahomotetrameric flavoprotein containing TPP, and it catalyzesthe oxidative decarboxylation of pyruvate to acetate and CO2,probably using a menaquinone as the physiological electronacceptor. Using 2,6-dichloroindophenol as an artificial elec-tron acceptor, the PQO activity was highest (about 0.055 U/mgof protein) in C. glutamicum cells grown on complex mediumand about threefold lower when glucose was added to thecomplex medium or when the cells were grown in minimalmedium containing different carbon sources (52). The PQOthus represents an additional pyruvate-decarboxylating en-

zyme in C. glutamicum, and we therefore speculated that thePQO reaction, together with the reactions of the acetate kinaseand the phosphotransacetylase, might be able to bypass thePDHC reaction (52). However, the inability of the PDHC-negative C. glutamicum mutant to grow on minimal mediumcontaining glucose indicated that the PQO is not able to com-pensate for a PDHC under the conditions generally used tocultivate C. glutamicum. The inability to replace the PDHCmight be due to the relatively low affinities of the PQO forpyruvate (Km � 30 mM [52]) and/or of the acetate kinase foracetate (Km � 7.9 mM [45]). However, in nature or underdifferent culture conditions there might be conditions when theintracellular pyruvate concentration in C. glutamicum rises tovalues well above the Km value, and then the PQO mightsubstitute for the PDHC.

Although the PDHC has a crucial role within the metabo-lism of C. glutamicum, relatively little effort has been devotedto the regulation of this enzyme complex on the genetic level.The analysis of the specific PDHC activities in extracts of cellsgrown on different media indicated that the synthesis or theassembly of the PDHC in C. glutamicum is only very weaklyregulated by the carbon source in the growth medium. Inaccordance with our findings, enzyme measurements with ex-tracts of C. glutamicum cells cultivated in a chemostat at var-ious dilution rates on minimal media containing glucose, pyru-vate, or lactate revealed no significant differences in thespecific activities under all conditions applied (8, 9, 64). Theseresults suggested that the PDHC is constitutively, and withabout the same specific activity, present in C. glutamicum cellsgrown in minimal medium containing different carbon sources.This is in contrast to the situation with E. coli, where two- tofourfold-higher activity and about-twofold-lower activity wasobserved in minimal medium containing pyruvate and acetate,respectively, than in minimal medium containing glucose (34,43, 44). However, under batch culture conditions, we found thespecific PDHC activity in C. glutamicum to be significantlydependent on the growth phase. The low PDHC activity at thebeginning and at the end of growth might be explained by anineffective or unbalanced assembly of the multienzyme com-plex or by low de novo synthesis of one or more enzymes of thePDHC in the respective growth phases. However, the tran-scriptional-fusion experiments showed no correlation betweenthe specific PDHC activity and the aceE promoter activity inthe course of the growth. Thus, it seems likely that the amountof functional PDHC in C. glutamicum depends on the expres-sion of the gene(s) encoding E2p and/or the LPD and, thus, onthe availability of these enzymes. However, this hypothesis andthe nature and mechanism of the genetic (co)control of theE1p, E2p, and the LPD genes in C. glutamicum still have to beelucidated.

Our compilation of E1 sequences deposited in public data-bases suggests that the distribution of homodimeric and het-erotetrameric E1p subunits in bacteria is not in accordancewith the rRNA-based phylogeny of bacteria and, instead, ismuch more heterogeneous than previously described (11, 17,37). The analyses indicate that either homodimeric or het-erotetrameric E1p enzymes are present in a given bacterialgroup; e.g., homodimeric enzymes are found mainly in �- and-proteobacteria, in high-GC-content gram-positive bacteria,and in deinococci, whereas heterotetrameric enzymes are

TABLE 4. Protein sequences identities among representativeheterotetrameric E1p �-subunits

Organisma

% Identity

Zymomonasmobilis

Bacillussubtilis

Zymomonas mobilis 33Rickettsia rickettsii 53 29Caulobacter crescentus 59 30Bradyrhizobium japonicum 59 30Agrobacterium tumefaciens 58 32Cytophaga hutchinsonii 47 28Mus musculus 49 26Rattus norvegicus 51 27Saccharomyces cerevisiae 49 28Acholeplasma laidlawii 33 42Listeria monocytogenes 28 74Oceanobacillus iheyensis 30 72Lactobacillus plantarum 29 53Bacillus subtilis 33Mycoplasma capricolum 29 32Geobacter sulfurreducens 28 44Coxiella burnetii 30 40Thermus thermophilus 33 42Haloferax volcanii 29 41Synechocystis sp. 39 27Prochlorococcus marinus 43 30Synechococcus elongatus 42 28Arabidopsis thaliana 43 29

a For accession numbers, see Materials and Methods.



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found predominantly in �-proteobacteria, cyanobacteria, andthe low-GC-content gram-positive bacteria. A simple explana-tion for this heterogeneous distribution of the two E1p types isthat homodimeric and heterotetrameric E1 progenitors haveexisted in ancestral bacteria and that later, in the process ofdiversification, the enzymes in different bacterial groups ac-quired specialized (e.g., pyruvate-specific) functions. This hy-

pothesis would explain (i) that homodimeric and heterotet-rameric E1p proteins show no obvious sequence similarity (11,17), (ii) that both E1 types possess completely different se-quence motifs important for catalysis (17), and (iii) that somebacteria (e.g., C. burnetii and T. thermophilus) (Fig. 6 and 7)possess genes for proteins with significant similarity to bothhomodimeric and heterotetrameric E1p enzymes.

FIG. 7. Phylogenetic tree of the heterotetrameric E1� protein family. Numbers at the nodes give the bootstrap values. Organisms are listed inMaterials and Methods. Arrows denote proposed gene duplications. Abbreviation: -bc, E1 �-subunit of a branched-chain 2-oxoacid dehydrogenasecomplex.



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Analysis of the phylogeny of the homodimeric E1p family(Fig. 6) reveals that the cluster comprising the enzymes of thehigh-GC-content gram-positive bacteria and deinococci alsocontains several enzymes of -proteobacteria, e.g., Legionellaand Acinetobacter (cluster II). Thus, those proteins are prob-ably more closely related to the proteins of the high-GC-con-tent gram-positive bacteria and deinococci than to those ofother -proteobacteria, e.g., E. coli or Haemophilus. Further-more, the homodimeric E1p proteins of the -proteobacteriaAzotobacter, Microbulbifer, and Pseudomonas are located in acompletely independent cluster (cluster IV in Fig. 6). Thisheterogeneity suggests that the enzymes may originate fromindependent gene transfer events from ancient bacteria intoancestral -proteobacteria. Further analysis of the phylogenyof the homodimeric E1p family revealed that the eukaryoticAnopheles gambiae protein is positioned within the -pro-teobacterial cluster (cluster I in Fig. 6) and that the three-proteobacterial Xanthomonas, Xylella, and Methylococcusproteins are located within the main �-proteobacterial clusterof enzymes (cluster III in Fig. 6). These findings probablyrepresent examples for more recent gene transfers from bac-teria to eukaryotes and from -proteobacteria to �-proteobac-teria.

The phylogeny of the heterotetrameric E1 family (Fig. 7)relates mitochondrial and chloroplast E1p �-subunits with E1p�-subunits of the �-proteobacteria and cyanobacteria, respec-tively, which is in agreement with endosymbiotic gene transferfrom bacteria to eukaryotes (7, 16, 51). Furthermore, the pu-tative heterotetrameric E1p �-subunit of the archaeonHaloferax is positioned within the cluster containing mainlyproteins of the low-GC-content gram-positive bacteria (clusterI in Fig. 7), suggesting that this E1p �-subunit gene has beenrecruited via gene transfer from the gram-positive bacteria.The obvious absence of PDHCs in other archaea (70) is inagreement with the finding that the oxidative decarboxylationof pyruvate is generally catalyzed by pyruvate:ferredoxin oxi-doreductases in archaea (31, 32, 47).

Interestingly, the phylogenetic analysis revealed that all het-erotetrameric E1p �-subunits of the low-GC-content gram-positive bacteria cluster with the known heterotetrameric E1�-subunits of the BCOADHC in many bacteria (Fig. 7, clusterI). This clustering indicates that the heterotetrameric E1p pro-teins of this lineage are more closely related to E1b proteinsthan to heterotetrameric E1p enzymes in �-proteobacteria andcyanobacteria. Thus, we conclude that it is likely that the E1pproteins of the low-GC-content gram-positive lineage are de-rived from E1b proteins of a BCOADHC via a relatively recentgene duplication event in an ancestral low-GC-content gram-positive bacterium. This hypothesis is corroborated by the find-ing that B. subtilis E1p catalyzes both the decarboxylation ofpyruvate (Km � 10.2 �M, Vmax � 60 mU/mg protein) and ofbranched-chain 2-oxoacids (Km � 33.6 to 88.8 �M, Vmax � 11.8to 53.0 mU/mg protein) (39). The cyanobacterial heterotet-rameric E1p �-subunits also are more closely related to E1b�-subunits than to the respective E1p �-subunits of �-pro-teobacteria, suggesting that the cyanobacterial heterotet-rameric E1p proteins also arose from E1b proteins via a lessrecent gene duplication event in an ancestral strain. The hy-pothesis that the heterotetrameric E1p proteins derived fromE1b proteins, rather than the reverse direction, is in agreement

with the assumption that early environments have been rich inamino acids and thus that amino acid-degrading enzymes suchas BCOADHCs should be an earlier development in the evolu-tion than the “aerobic” PDHCs. The notion that BCOADHCshave existed early in evolution is substantiated by the fact thatBCOADHCs are present in archaea, which indicates thatBCOADHC have developed before the divergence of bacteriaand archaea (22). The finding that heterotetrameric PDHCsare abundant only in �-proteobacteria, low-GC-content gram-positive bacteria, and cyanobacteria thus suggests that het-erotetrameric PDHCs trace back to developments that prob-ably occurred late during the diversification of bacteria. Inconclusion, we propose that heterotetrameric E1p subunits inbacteria arose from ancient E1b proteins via multiple indepen-dent gene duplication events.

ACKNOWLEDGMENTS

We thank Joy Schreiner for critically reading the manuscript.The support of the EC (grant VALPAN, QLK3-2000-00497), the

Fachagentur Nachwachsende Rohstoffe of the BMVEL (grant04NR004/22000404), and the Grant Agency of the Czech Republic(grant 525/04/0548) is gratefully acknowledged.

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