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Journal of Bacteriology, May 2002, p. 2728-2739, Vol. 184, No. 10
0021-9193/02/$04.00+0     DOI: 10.1128/JB.184.10.2728-2739.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.

Identification of Two prpDBC Gene Clusters in Corynebacterium glutamicum and Their Involvement in Propionate Degradation via the 2-Methylcitrate Cycle

Wilfried A. Claes, Alfred Pühler, and Jörn Kalinowski*

Lehrstuhl für Genetik, Fakultät für Biologie, Universität Bielefeld, 33501 Bielefeld, Germany

Received 10 December 2001/ Accepted 22 February 2002


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Genome sequencing revealed that the Corynebacterium glutamicum genome contained, besides gltA, two additional citrate synthase homologous genes (prpC) located in two different prpDBC gene clusters, which were designated prpD1B1C1 and prpD2B2C2. The coding regions of the two gene clusters as well as the predicted gene products showed sequence identities of about 70 to 80%. Significant sequence similarities were found also to the prpBCDE operons of Escherichia coli and Salmonella enterica, which are known to encode enzymes of the propionate-degrading 2-methylcitrate pathway. Homologous and heterologous overexpression of the C. glutamicum prpC1 and prpC2 genes revealed that their gene products were active as citrate synthases and 2-methylcitrate synthases. Growth tests showed that C. glutamicum used propionate as a single or partial carbon source, although the beginning of the exponential growth phase was strongly delayed by propionate for up to 7 days. Compared to growth on acetate, the specific 2-methylcitrate synthase activity increased about 50-fold when propionate was provided as the sole carbon source, suggesting that in C. glutamicum the oxidation of propionate to pyruvate occurred via the 2-methylcitrate pathway. Additionally, two-dimensional gel electrophoresis experiments combined with mass spectrometry showed strong induction of the expression of the C. glutamicum prpD2B2C2 genes by propionate as an additional carbon source. Mutational analyses revealed that only the prpD2B2C2 genes were essential for the growth of C. glutamicum on propionate as a sole carbon source, while the function of the prpD1B1C1 genes remains obscure.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
The gram-positive, nonsporulating soil microorganism Corynebacterium glutamicum is widely used in biotechnological processes for the fermentative production of amino acids, nucleotides, and other metabolites (60). Up to now, a variety of genes primarily involved in amino acid production were characterized at the molecular level, and subsequently, the synthesis of selected amino acids was improved by genetic engineering of the biosynthetic pathways of this organism (29). Additionally, knowledge of central metabolic pathways in C. glutamicum is of growing interest for the deduction of metabolic fluxes in this organism (9, 11, 40). One of the most important pathways in the central metabolism is the citric acid cycle, which provides several metabolic precursors for amino acid production. Due to the observation that a C. glutamicum strain with reduced citrate synthase activity produced significant amounts of aspartate and lysine (43), the C. glutamicum citrate synthase gene gltA (EC 4.1.3.7) was cloned and sequenced (12). A C. glutamicum mutant strain with a disrupted gltA gene revealed a glutamate auxotrophy and the loss of detectable citrate synthase activity on all carbon sources tested (12). On the basis of these findings and additional hybridization experiments, it was concluded that gltA is the only citrate synthase gene in C. glutamicum.

A reaction similar to the one catalyzed by citrate synthases is observed in the 2-methylcitrate cycle, in which propionate is {alpha}-oxidized to pyruvate. This pathway was originally identified for Yarrowia lipolytica (50) and shown to be present in several filamentous fungi (32) and yeast (38). Recently, various experiments have shown the existence of the 2-methylcitrate cycle in the enterobacteria Escherichia coli and Salmonella enterica serovar Typhimurium (22, 30, 55). The genes for propionate breakdown in these organisms are located in two adjacent transcription units, prpR and prpBCDE, and are, with the exception of prpE, essential for growth on propionate as the sole carbon and energy source (21). While the prpBCDE genes encode the enzymes of the 2-methylcitrate cycle in S. enterica (22), prpR is the transcriptional activator of this gene cluster (36). A prp locus displaying a different gene arrangement was very recently identified in the gram-negative Ralstonia eutropha (4).

In S. enterica, activation of propionate to propionyl-coenzyme A (CoA) is mediated by the propionyl-CoA synthetase PrpE (23) in an ATP-dependent manner. The following condensation reaction of propionyl-CoA and oxaloacetate to 2-methylcitrate is catalyzed by the 2-methylcitrate synthase PrpC (EC 4.1.3.31), and the catalytic mechanism of the 2-methylcitrate synthase reaction is presumed to be identical to that for the citrate synthase reaction (14). The isomerization of 2-methylcitrate to 2-methylisocitrate with 2-methyl-cis-aconitate as an intermediate requires the action of the 2-methylcitrate dehydratase (51). In S. enterica, the PrpD protein (EC 4.2.1.79) catalyzes this step, whereas the aconitase AcnA or AcnB is responsible for the subsequent hydratase reaction (24). Finally, the resulting 2-methylisocitrate is cleaved by the prpB gene product (EC 4.1.3.30) to pyruvate and succinate (6, 49). While oxaloacetate is regenerated from the latter by the citric acid cycle, pyruvate may be further oxidized to acetyl-CoA or may serve directly as a building block for biosyntheses.

Although several pathways for propionate degradation have been proposed (reviewed in reference 55), it is now assumed—mainly based on genome data—that the 2-methylcitrate cycle may be widespread among bacteria (22) and may represent the major pathway for propionate oxidation under aerobic conditions (7). However, until now no experimental data have been published for the operation of the 2-methylcitrate cycle in gram-positive bacteria.

In the present paper, we report on the identification, cloning, and molecular characterization of two citrate synthase homologous genes in C. glutamicum, which are located in two distinct prpDBC gene clusters. The results concerning overexpression of the C. glutamicum prpC1 and prpC2 genes, analyses of crude C. glutamicum protein extracts after growth on propionate, and mutational analyses of both prpDBC gene clusters were used to address the question of which gene cluster is responsible in C. glutamicum for the degradation of propionate.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Bacterial strains, plasmids, and growth conditions. Bacterial strains and plasmids used in this study are listed in Table 1. E. coli and C. glutamicum strains were routinely grown in Luria-Bertani (LB) medium (41) with 10 mM glucose at 37 and 30°C, respectively. Liquid medium CGXII (28) and solid medium MM1, described previously as MMYE (27) without yeast extract, were used for growth of C. glutamicum on minimal media. Carbon and energy sources for growth on minimal media were used as indicated. Antibiotics for plasmid selection were kanamycin (50 µg ml-1 for E. coli and 25 µg ml-1 for C. glutamicum) and ampicillin (100 µg ml-1 for E. coli).


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  		TABLE 1. Bacterial strains and plasmids used in this study

 
DNA isolation, manipulation, and transfer. E. coli DH5{alpha}MCR was used for routine recombinant DNA experiments. Plasmid DNA of E. coli was prepared by the alkaline lysis technique (41), modified for C. glutamicum by using 20 mg of lysozyme ml-1 in buffer HB1 at 37°C for 2 h. Plasmid DNA for sequencing was isolated by means of the Qiagen Plasmid Spin Prep Kit according to the manufacturer's protocol (Qiagen, Hilden, Germany). Total chromosomal DNA of C. glutamicum was prepared according to the method of Tauch et al. (52). DNA restriction, agarose gel electrophoresis, Klenow treatment, and ligation were carried out as described elsewhere (41). DNA restriction fragments required for cloning were purified from agarose gels by using the Nucleotrap Extraction Kit for Nucleic Acids (Macherey-Nagel, Düren, Germany). E. coli and C. glutamicum were transformed by electroporation (18, 54) using the Bio-Rad Gene Pulser system (Bio-Rad, Munich, Germany). Cloning of PCR fragments and introduction of resulting plasmids into E. coli TOP10 was performed using the Zero Blunt TOPO PCR Cloning Kit from Invitrogen (San Diego, Calif.).

PCR techniques. PCR experiments were carried out with a PTC-100 thermocycler from MJ Research Inc. (Watertown, Mass.) with proof-reading Pfu DNA polymerase (Stratagene, La Jolla, Calif.). Initial denaturation was conducted at 94°C for 2 min followed by denaturation for 45 s, annealing for 45 s at primer-dependent temperature at Tm (2AT + 4GC) - 5°C (48) and extension at 72°C for 90 s. This cycle was repeated 35 times, followed by a final extension step for 8 min at 72°C. PCR products were purified using the PCR Purification Spin kit (Qiagen).

Southern hybridization. Chromosomal DNA of C. glutamicum was digested with restriction endonucleases, separated on 0.8% agarose gels, and blotted onto Hybond-N nylon membranes (Amersham Pharmacia Biotech, Freiburg, Germany) using the vacuum blotting system VacuGene XL (Amersham Pharmacia Biotech). Fixation of DNA, labeling of DNA probes, hybridization, and detection were performed with the DIG DNA Labeling and Detection Kit Nonradioactive from Roche (Mannheim, Germany).

DNA sequencing and sequence analysis. DNA sequences of cosmids cg2m19 and cg6d10 were determined as described elsewhere (53). Plasmid DNA inserts were sequenced by the DNA sequencing group of the IIT Biotech GmbH (Bielefeld, Germany). DNA sequence data were assembled and processed by means of the computer programs of Staden (46). For sequence interpretation during the progress of the C. glutamicum genome project, the automated sequence investigation programs MAGPIE (13) and GenDB (University of Bielefeld) were used. Searches for DNA and amino acid sequence similarities were carried out with the BLAST program package (1). Global multiple alignments between protein sequences were calculated with the CLUSTAL W program (56). Energies of RNA secondary structures were predicted with MFOLD (61).

Construction of deletion mutant strains of C. glutamicum RES167. Defined chromosomal deletions within the gltA gene and within the prpD1B1C1 and prpD2B2C2 gene regions of C. glutamicum RES167 were constructed using the sacB system, which helps to identify an allelic exchange by homologous recombination (42). The positions of deleted nucleotides for each gene and the corresponding amino acids are given in Table 1.

Deletion mutations within open reading frames (ORFs) of the prpD1B1C1 gene cluster (C. glutamicum strains WAC1 to WAC4) were introduced by using unique restriction sites in the respective genes (Table 1). Southern hybridization experiments using EcoRI-SacI-digested chromosomal DNAs and digoxigenin (DIG)-labeled plasmid pCLA1 were performed to verify the construction of the chromosomal prpD1B1C1 deletions.

Deletions within ORFs of the prpD2B2C2 gene cluster (C. glutamicum strains WAC5 to WAC8) were introduced by gene "SOEing," following the method of Horton (25). Therefore, the DNA regions upstream and downstream of the gene to be deleted were amplified by two PCRs. Cosmid cg6d10 was used as template DNA, and PCR primer pairs were designed to give a fragment length of about 1 kb. For the amplification of the upstream DNA region, each inner PCR primer flanking the deletion carried a 20-nucleotide (nt) linker, whereas each inner PCR primer flanking the deletion at the downstream region carried the reverse complement linker. Thus, both PCR products from the initial two PCRs overlap over the introduced 20-bp linker with the sequence 5'-TCGTCCATTTAAATCCTGCT-3'. Purified PCR product (0.5 µl) from the initial two reactions was mixed, and the mixture served as template DNA for the third PCR. Herein, the outer primers of the initial two reactions were used to amplify the deletion fragment. The resulting 2-kb fragment carrying the deletion with the central 20-bp linker DNA was cloned into vector pCR-Blunt-II-TOPO and subsequently subcloned as a SpeI-XhoI fragment into the XbaI-SalI-digested vector pK18mobsacB. The DNA sequence of each of the constructed deletion alleles was verified by sequencing before introduction of the deletion into the C. glutamicum chromosome. Southern hybridization experiments with the DIG-labeled plasmid pCLA5 and KpnI-DraI-digested genomic DNA from C. glutamicum were performed to confirm the introduced deletions in the prpD2B2C2 locus.

To delete the gltA gene, the plasmid pUC18-gltA was digested with EcoRV and NruI and subsequently ligated, thereby deleting an internal 1,286-bp fragment of the gltA coding region, corresponding to amino acid residues 6 to 434. The deletion fragment of the resulting plasmid was subcloned as a SalI-HindIII fragment into the vector pK18mobsacB. Southern hybridization experiments with DIG-labeled pUC18-gltA and HindIII-SalI-digested genomic DNA from prescreened glutamate-auxotrophic C. glutamicum strains confirmed the introduced gltA deletion. The resulting strain was designated C. glutamicum GLTA1.

Construction of prpC1 and prpC2 expression plasmids. The C. glutamicum prpC1 gene was cloned with its own putative ribosome binding site as a SacI-XhoI fragment (nt 4,576 to 5,937) from pCLA1 into SacI-SalI-digested vector pTrc99a, resulting in the plasmid pTrc-prpC1. For homologous expression, the prpC1 gene was subcloned from the plasmid pTrc-prpC1 as an EcoRI-MunI fragment and ligated into the EcoRI-digested C. glutamicum expression vector pZ8-1, resulting in the plasmid pZ-prpC1.

The prpC2 gene was amplified with its own ribosome binding site as a 1,266-bp DNA fragment (nt 3,861 to 5,128) by PCR with Pfu DNA polymerase and primers C2.1 (5'-ATTCCGCTGACAGCTACAAG-3') and C2.2 (5'-GAATGTTGCACCATGGCT-3'). The purified PCR product was ligated into the vector pCR-Blunt-II, and the resulting vector was subsequently digested with EcoRI. The prpC2-containing fragment was isolated from an 0.8% agarose gel and cloned into EcoRI-digested pTrc99a or pZ8-1, respectively, to give the plasmids pTrc-prpC2 and pZ-prpC2. The sequence of the amplified prpC2 gene and the correct orientation of the cloned fragment were validated by sequencing. Induction of prpC1 and prpC2 expression in E. coli DH5{alpha}MCR on pTrc99a-based plasmids was carried out by addition of 0.5 mM isopropyl-ß-D-thiogalactopyranoside (IPTG) to LB-grown cultures at an optical density at 580 nm of 0.4 and growth for an additional 4 h.

Preparation of crude protein extracts of bacterial cultures. E. coli and C. glutamicum protein extracts were prepared from cells grown to the late exponential growth phase in liquid medium CGXII or LB. Forty-five milliliters of bacterial culture was harvested by 10 min of centrifugation at 5,500 x g. Cell pellets were washed twice in 30 ml of 50 mM Tris-HCl (pH 7.5) and finally centrifuged for 10 min at 5,500 x g. After removal of the supernatant, cells were resuspended in 1 ml of 50 mM Tris-HCl (pH 7.5) containing 10 µg of RNase A and 1 µg of DNase I. One milliliter of the cell suspension was added to a RiboLyser BLUE tube (Hybaid, Ltd., Teddington, United Kingdom) containing a silica-ceramic matrix. Cell disruption using the Hybaid RiboLyser was carried out at a speed ratio of 6.5 for two time intervals of 30 s. Cell debris was removed by 30 min of centrifugation at 14,900 x g. Protein concentrations of the crude extracts were determined by means of the Bio-Rad Protein Assay with bovine serum albumin as reference. Cell extracts were used immediately or frozen in aliquots at -80°C.

Proteome analysis. For 2-dimensional polyacrylamide gel electrophoresis (2D-PAGE [34]) of proteins, 400 µl of crude protein extract was precipitated for 4 h at -20°C with 1.6 ml of acetone and subsequently centrifuged at 14,900 x g for 45 min. Dry protein pellets were resuspended for 1 h in 400 µl of rehydration buffer, consisting of 9 M urea, 2% (wt/vol) 3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate, 0.5% (vol/vol) IPG buffer (Amersham Pharmacia Biotech), 0.28% (wt/vol) dithiothreitol (DTT), and a few grains of bromphenol blue. For isoelectric focusing (IEF), precast IPG gels with a linear pH gradient of 4 to 7 and an IPGphor IEF unit (Amersham Pharmacia Biotech) were used. Proteins were focused for 28 h in six steps, 2 h at 0 V, 6 h at 30 V, 6 h at 60 V, 1 h at 500 V, 1 h at 1,000 V, and 12 h at 8,000 V. Focused IPG gels were equilibrated twice for 15 min in a buffer containing 50 mM Tris-HCl (pH 8.8), 6 M urea, 30% (vol/vol) glycerin, 2% (wt/vol) sodium dodecyl sulfate, and 1% (wt/vol) DTT. For the second equilibration step, DTT was replaced by 4.5% (wt/vol) iodoacetamide. The second dimension was run using precast ExcelGel Gradient XL 12 to 14% gels (Amersham Pharmacia Biotech) on a Multiphor II apparatus (Amersham Pharmacia Biotech) as recommended by the manufacturer, and gels were subsequently stained with Coomassie brilliant blue (41). Protein spots were excised from gels and digested with modified trypsin (Promega, Mannheim, Germany), and peptide mass fingerprints were determined with a Biflex III mass spectrometer (Bruker Daltonics, Bremen, Germany) and analyzed with the MASCOT software (37), as described elsewhere (19).

Enzyme assays. Citrate synthase and 2-methylcitrate synthase activities were assayed spectrophotometrically with an LKB Biochrom 4060 photometer (Amersham Pharmacia Biotech) at 30°C according to the method of Srere (45). One milliliter of reaction mixture contained 50 mM Tris-HCl (pH 8.0), 1 mM 5,5'-dithiobis-(2-nitrobenzoate), 50 µl of crude protein extract, adjusted by dilution in 50 mM Tris-HCl (pH 7.5) to contain up to 0.5 U of enzyme ml-1, and 0.3 mM acetyl-CoA or propionyl-CoA, respectively. After measurement of unspecific acyl-CoA deacylase activity by absorption (A412) for 2 min, the reaction was started by addition of 50 µl of 0.5 mM oxaloacetate, monitoring the A412 for an additional 3 min.

Nucleotide sequence accession numbers. The nucleotide sequences of the prp gene loci of C. glutamicum were deposited in the GenBank database under the accession numbers AF434798 and AF434799.


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
DNA sequence analysis revealed the existence of two different prpDBC gene clusters in Corynebacterium glutamicum ATCC 13032. The C. glutamicum citrate synthase gene gltA was previously described by Eikmanns and colleagues (12). In the meantime, the C. glutamicum genome was sequenced by using cosmid and bacterial artificial chromosome libraries (20, 53). The sequences obtained were annotated by the automated genome interpretation programs MAGPIE (13) and GenDB (University of Bielefeld) and checked for further genes homologous to the gltA sequence. Two candidate genes were found, one located on cosmid cg2m19 and the other on cosmid cg6d10. Through inspection of the surroundings of the genes identified, it became evident that they belong to larger gene regions consisting of three genes (Fig. 1) characterized by significant identities to each other and also to the prp gene regions encoding proteins of the 2-methylcitrate cycle of S. enterica, E. coli and R. eutropha (4, 21, 55). The two gene clusters of C. glutamicum were therefore termed prpD1B1C1 and prpD2B2C2. For further analysis, the prpD1B1C1 gene cluster was subcloned from cosmid cg2m19 as a native 6.4-kb EcoRI DNA fragment into vector pUC19, resulting in the plasmid pCLA1. The prpD2B2C2 region was cloned from cosmid cg6d10 as a native 6.0-kb KpnI DNA fragment into the vector pMECA, resulting in the plasmid pCLA5 (Fig. 1).



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  		FIG. 1. Physical and genetic maps of different bacterial prp gene regions. (a and b) Restriction maps of the C. glutamicum prpD1B1C1 and prpD2B2C2 gene fragments, subcloned DNA fragments, and deletion mutants. (c) Genetic organizations of the prp gene clusters of E. coli (GenBank accession no. U73857), S. enterica serovar Typhimurium (GenBank accession no. U51879), and R. eutropha (GenBank accession no. AF325554 and AF331923). Only the restriction sites used for cloning of both C. glutamicum prp loci are shown. The positions of the C. glutamicum ORFs and genes are indicated by arrows, and putative rho-independent termination structures are presented as hairpin structures. The deletions introduced into the C. glutamicum chromosome are described in detail in the Material and Methods section. Homologous genes within the prp regions of E. coli, S. enterica, R. eutropha, and C. glutamicum are shaded grey.

 
Computational analyses showed that the two C. glutamicum prpDBC loci are highly similar to each other with respect to their genes and the corresponding gene products (Table 2). The sequence identity averages between 70 and 80% on the nucleotide level as well as on the amino acid level. Thus, it is assumed that both prpDBC regions of C. glutamicum are paralogous, originating by gene duplication and diversification. Compared to the C. glutamicum gltA gene product, which consists of 437 amino acids with a molecular mass of 48.9 kDa (12), both prpC gene products are different in size and amino acid sequence. The sequence identities of PrpC1 and PrpC2 to GltA on the amino acid level are 32 and 33%, respectively.


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  		TABLE 2. Comparison of C. glutamicum prp genes and gene products

 
The amino acid sequence identities of the C. glutamicum prp gene products to the respective proteins of S. enterica comprised about 26% for PrpD1 and PrpD2, 42% for PrpB1 and PrpB2, and 40% for PrpC1 and PrpC2 (Table 2). Commensurate sequence similarities were found to the Prp proteins of the gram-negative bacteria E. coli and R. eutropha (data not shown). Amino acid sequence identities of about 25% to recently identified eukaryotic 2-methylcitrate cycle enzymes (7, 31) were found for the C. glutamicum PrpB1 and PrpB2 proteins and for PrpC1 and PrpC2. Due to these similarities, prpC1 and prpC2 are supposed to code for 2-methylcitrate synthases, prpB1 and prpB2 for 2-methylisocitrate lyases, and prpD1 and prpD2 for 2-methylcitrate dehydratases.

The C. glutamicum PrpD proteins shared no sequence similarities to other proteins with known function but revealed homologies to putative ORFs of gram-positive bacteria. The designation of C. glutamicum PrpB1 and PrpB2 as 2-methylisocitrate lyases is based—in addition to the homology data—on the identification of the amino acid sequence KRCGH in both proteins (data not shown). On the basis of this motif, bacterial 2-methylisocitrate lyases may become distinguishable from isocitrate lyases, since a KKCGH amino acid motif is supposed to represent the catalytic site of the latter group of enzymes (6). Conserved amino acid residues essential for citrate synthase activity were found in both deduced C. glutamicum prpC gene products as well as in all known 2-methylcitrate synthases (data not shown), thus indicating similar reaction mechanisms for the two enzymes (14).

Although the two C. glutamicum prp loci were highly similar (Table 2), the flanking regions of both prpDBC gene regions of C. glutamicum shared no further common sequence properties. Furthermore, they revealed no homologies to genes known to be involved in propionate degradation, e.g., ORF5 of R. eutropha (4), or to those corresponding to aconitase-like enzymes, acting as 2-methyl-cis-aconitic acid hydratases, which were found in hypothetical prp loci of numerous bacterial genomes (4, 24). In contrast to the situation with S. enterica and E. coli (Fig. 1), neither the prpD1B1C1 nor prpD2B2C2 gene cluster of C. glutamicum is in proximity to a prpE-homologous gene encoding propionyl-CoA synthetase.

The two prp loci showed an overall G+C content of 56.6 mol% (prpD1B1C1) and 55.5 mol% (prpD2B2C2), which agreed with the range of 55 to 57.7 mol% reported for the C. glutamicum chromosome (26). The prpD2B2C2 and prpD1B1C1 gene clusters were located in tandem orientation on the C. glutamicum chromosome and separated by 37,979 bp. Each may constitute an operon, since no transcriptional termination structures were found to be located between the prpD, prpB, and prpC coding regions of both loci. Moreover, the coding region of the prpB1 gene overlaps at its 3' end the prpC1 gene for 8 nt, and the 5' end of prpB1 overlaps the prpD1 3' end over 4 nt. Thus, it seemed possible that prpD1, prpB1, and prpC1 constitute an operon and that these genes are translationally coupled, since overlapping of the stop codon and the start codon plays a role in translational reinitiation in E. coli (35). In contrast to the first gene cluster, only the genes prpD2 and prpB2 were found to be possibly translationally coupled, since their stop and start codons overlap for 1 nt, whereas the coding regions of prpB2 and prpC2 were separated by 28 bp. Putative ribosome binding sites matching the optimal C. glutamicum ribosome binding site with the sequence 5'-AGAAAGGAGG-3' (2) were located in front of each of the prp genes of both loci (Table 2) in a distance of 4 to 13 nt.

An imperfect inverted repeat of 47 bp could be located 33 bp downstream of the C. glutamicum prpC1 gene, which displayed a {Delta}G (30°C) of -111.0 kJ as calculated with MFOLD (61). The palindromic sequence was followed by a stretch of four T-residues, and therefore this region may act as a rho-independent transcriptional termination signal. For the prpD2B2C2 gene cluster, two putative rho-independent transcriptional terminators with a {Delta}G (30°C) of -62.4 kJ mol-1 and of -121.5 kJ mol-1 were identified 77 and 102 bp downstream of the prpC2 gene, respectively. Summarizing these sequence data, it can be stated that both C. glutamicum prpDBC regions seem to be potentially responsible for encoding enzymes of the 2-methylcitrate cycle.

The C. glutamicum prpC1 and prpC2 genes encode enzymes with activities as citrate synthases and 2-methylcitrate synthases. Since the deduced C. glutamicum prpD1B1C1 and prpD2B2C2 gene products exhibited strong amino acid sequence similarities to enzymes of the 2-methylcitrate cycle, we addressed the question of whether the prpC1 and prpC2 gene products act as 2-methylcitrate synthases, the key enzyme group within the 2-methylcitrate cycle. In order to test the ability of both PrpC proteins to catalyze the condensation reaction from propionyl-CoA and oxaloacetate to 2-methylcitrate, both prpC genes were subcloned with their endogenous putative ribosome binding sites into the C. glutamicum expression vector pZ8-1 equipped with the constitutive tac promoter. The resulting plasmids, pZ-prpC1 and pZ-prpC2, and also the control vector, pZ8-1, were used to transform the C. glutamicum strain GLTA1, in which the gltA gene was deleted. Like the previously described C. glutamicum gltA insertion mutant (12), strain GLTA1 was devoid of detectable citrate synthase activity and unable to grow on solid minimal medium unless supplemented with 5 mM glutamate.

After growth to late exponential growth phase in rich medium LB, crude protein extracts of C. glutamicum GLTA1 harboring either plasmid pZ-prpC1 or pZ-prpC2 showed an additional protein band of about 42 kDa after one-dimensional sodium dodecyl sulfate-PAGE analysis, thus indicating the strong expression of both prpC genes in the homologous system (data not shown). Tryptic digests and matrix-assisted laser desorption ionization-mass spectrometry fingerprints of these protein bands verified this assumption. Furthermore, the glutamate-auxotrophic phenotype of the C. glutamicum GLTA1 strain during growth on minimal medium MM1 was complemented by both the pZ-prpC1 and pZ -prpC2 plasmids but not by the control vector pZ8-1, which indicated a function of both PrpC proteins as citrate synthases in C. glutamicum.

To confirm the complementation experiments, enzyme activity tests were performed for citrate synthase and 2-methylcitrate synthase activity (Table 3). In the citrate synthase-negative strain C. glutamicum GLTA1 transformed with the control vector pZ8-1, no citrate synthase activity was detectable after growth on rich medium, which agrees with the findings of Eikmanns and colleagues (12). Additionally, no significant 2-methylcitrate synthase activity was observed. In contrast to the control strain, the strains harboring the expression vector pZ-prpC1 or pZ-prpC2 revealed significant increases in both citrate synthase and 2-methylcitrate synthase activities (Table 3). Thus, the prpC1 and prpC2 gene products catalyzed the condensation reaction of oxaloacetate with both CoA derivatives, acetyl-CoA (the citrate synthase reaction) and propionyl-CoA (the 2-methylcitrate synthase reaction). Thereby, the specific activities for the different CoA substrates were different for the two C. glutamicum PrpC proteins under overexpression conditions. PrpC1 revealed a 2.4-fold higher specific activity with acetyl-CoA than with propionyl-CoA as the substrate, whereas PrpC2 showed a 2.1-fold-higher specific 2-methylcitrate synthase activity than citrate synthase activity.


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  		TABLE 3. Specific enzyme activities of PrpC1 and PrpC2 as citrate synthases and 2-methylcitrate synthases

 
Supplementary to the expression studies in the homologous system, we cloned the C. glutamicum prpC genes into the IPTG-inducible E. coli expression vector pTrc99a. E. coli DH5{alpha} was transformed either with the control vector pTrc99a or with the resulting plasmids pTrc-prpC1 and pTrc-prpC2. Crude protein extracts of pTrc-prpC1 or pTrc-prpC2 harboring E. coli cells showed the heterologous expression of both C. glutamicum genes after IPTG treatment (data not shown). Strong increases in citrate synthase and 2-methylcitrate synthase activities could be observed (Table 3). The calculated ratios of 2-methylcitrate synthase activity to citrate synthase activity (MCS/CS) were similar to those calculated for the expression in the homologous system. Low levels of enzyme activities of strains carrying pTrc-prpC1 or pTrc-prpC2 in the absence of IPTG were a result of incomplete repression of the tac promoter (3).

Due to their genomic organization within both prp loci, the C. glutamicum prpC genes were classified as 2-methylcitrate synthase genes, even though PrpC1 apparently revealed a citrate synthase activity that was higher than its 2-methylcitrate synthase activity.

Growth of C. glutamicum on propionate resulted in the induction of a specific 2-methylcitrate synthase activity. In order to test whether C. glutamicum is able to utilize propionate as a carbon source, minimal medium CGXII supplemented with either glucose, acetate, or propionate or combinations of these carbon sources was inoculated with C. glutamicum RES167. Since growth on glucose or acetate started 12 to 18 h after inoculation, growth on propionate as the single carbon source showed a lag phase of 5 to 7 days but subsequently yielded similar growth rates, like those on acetate as the single carbon source (Fig. 2). Furthermore, propionate as an additional substrate delayed growth on glucose by 36 h, probably due to the toxic properties of propionate. In contrast, growth on acetate was only weakly delayed by the addition of propionate. A similar observation was previously described for the fungus Aspergillus nidulans (7), in which growth on acetate was not influenced by the addition of propionate. During growth of C. glutamicum on substrate mixtures of glucose and propionate, no diauxic shift was observed. This is in accordance to the observation that co-utilization of mixtures of glucose and acetate as sole carbon sources leads to monophasic growth in C. glutamicum (59).



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  		FIG. 2. Growth of C. glutamicum RES167 on propionate as the sole or additional carbon source. An LB overnight culture of C. glutamicum RES167 was washed twice in 50 mM Tris-HCl (pH 7.5), and 2 x 107 cells were used as inoculum for 60 ml of the minimal medium CGXII. Each carbon source was added with a final concentration of 4 g liter of medium-1. Cultivations were done in 250-ml Erlenmeyer flasks at 30°C and 150 rpm, and aliquots for the determination of the optical density at 580 nm were taken at the time intervals indicated.

 
Since PrpC1 and PrpC2 comprised activities as both citrate synthases and 2-methylcitrate synthases, we addressed the question of whether 2-methylcitrate synthase activity is induced during growth of C. glutamicum on propionate as the carbon source, thus giving clear evidence for the operation of the 2-methylcitrate cycle in C. glutamicum. For this purpose, C. glutamicum was grown in liquid medium CGXII with different carbon sources to late exponential growth phase, before enzyme activity tests were performed. Whereas only low levels of 2-methylcitrate synthase activity were detected during growth on glucose or acetate, enzyme activity was increased 14-fold (glucose) or 22-fold (acetate) when propionate was used as an additional carbon source (Table 4). The highest increase in 2-methylcitrate synthase activity (about 50-fold) was found during growth on propionate as the sole carbon source, and due to this strong induction of the 2-methylcitrate synthase activity it seems likely that propionate is metabolized through the 2-methylcitrate pathway in C. glutamicum.


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  		TABLE 4. Specific 2-methylcitrate synthase activities of C. glutamicum RES167 after growth on propionate

 
Parallel to the increase of 2-methylcitrate synthase activity, a 1.3- to 6-fold increase in citrate synthase activity was measurable after addition of propionate. The total specific citrate synthase activity amounts to circa 40% in comparison to the 2-methylcitrate synthase activity, e.g., during growth on propionate, and similar ratios were obtained when propionate was used as an additional carbon source. These results may indicate that PrpC2 supplied the main part of the measured 2-methylcitrate synthase activity during growth on propionate, since the overexpression experiments of prpC1 led to higher citrate than 2-methylcitrate synthase activities.

The gene products of the prpD2B2C2 locus were strongly induced during growth of C. glutamicum on propionate as an additional carbon source. To analyze the propionate-dependent expression of all prp genes, crude protein extracts were prepared after growth of C. glutamicum RES167 to the late exponential growth phase on minimal medium CGXII with acetate or with acetate plus propionate as carbon sources. For the identification of propionate-specific proteins, acetate was used as the control carbon source. Proteins were separated by 2D-PAGE and stained with Coomassie brilliant blue. With propionate as an additional carbon source, synthesis of six strongly increased or newly synthesized protein spots occurred (Fig. 3). Although the corresponding gene products of the two C. glutamicum prpDBC gene clusters were very similar with respect to their molecular masses and their isoelectric points (Table 2), a tryptic digest of the proteins in silico showed that they are clearly distinguishable by mass spectrometry. Therefore, propionate-induced protein spots were cut from the gel and digested with trypsin, and peptides were analyzed in a matrix-assisted laser desorption ionization-mass spectrometry fingerprint approach. Interestingly, all propionate-specific protein spots were gene products of the prpD2B2C2 cluster. The tryptic fingerprint revealed sequence coverages of at least 55% for each of the proteins, therefore unambiguously identifying them (data not shown). No protein spots specified by the prpD1B1C1 gene cluster could be observed under the conditions tested. PrpD2, PrpB2, and PrpC2 were represented in each case by two distinct spots, an observation which may indicate posttranslational modifications of these proteins.



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  		FIG. 3. 2D-PAGE of protein extracts of C. glutamicum during growth on acetate with or without propionate. Crude protein extracts of C. glutamicum RES167 were harvested in the late exponential growth phase after growth on minimal medium CGXII containing 4 g of acetate liter-1 (a) or 4 g of acetate plus 4 g propionate liter-1 (b) as sole carbon sources. Two hundred fifty micrograms of precipitated and resuspended proteins were subjected to 2D-PAGE. The pH range and the molecular masses of marker proteins are indicated on the gels. Areas of prpD2B2C2 protein spots of both gels are magnified for comparison.

 
The prpD2B2C2 gene cluster is essential for C. glutamicum during growth on propionate. Since we showed that PrpC1 and PrpC2 mediate enzymatic activities as both citrate synthases and 2-methylcitrate synthases, and since a specific 2-methylcitrate synthase activity occurred in cell extracts of C. glutamicum during growth on propionate as a single or additional carbon source, we questioned whether either of the two sequenced prp loci of C. glutamicum can support growth on propionate as the sole carbon source. For this purpose, the C. glutamicum mutant strains WAC1 to WAC4, carrying deletions within the prpD1B1C1 gene region, and mutants WAC5 to WAC8, carrying deletions in the prpD2B2C2 locus, were constructed (Fig. 1). To rule out functional complementation of single mutants by the paralogous genes of the other prp cluster, double mutants WAC9 to WAC12 carrying deletions in corresponding genes of both clusters were constructed (Table 1). The introduced deletion mutations were verified by Southern hybridization with restriction enzyme-digested genomic DNA of the mutant strains and DIG-labeled plasmids pCLA1 and pCLA5 (data not shown). The constructed C. glutamicum strains were used to test their abilities to grow on different carbon sources.

The constructed mutants showed no phenotypic alteration relative to the control strain, RES167, when grown either on rich medium or on minimal medium with glucose or acetate as the carbon source (data not shown). And similar to the observations with liquid medium, initial growth of the control strain, C. glutamicum RES167, on solid minimal medium with propionate as the sole carbon source was delayed by 10 to 14 days. The C. glutamicum strains WAC1 to WAC4, carrying deletions within the prpD1B1C1 gene cluster, were not affected in growth on propionate (Fig. 4) but showed the same delay in detectable growth as the control strain, RES167. In contrast, none of the mutants carrying deletions within the prpD2B2C2 cluster (strains WAC5 to WAC8) was able to grow on propionate as the sole carbon and energy source, even though the incubation time was prolonged for an additional week (Fig. 4). This implies that the prpD1B1C1 genes were apparently not able to complement the loss of the prpD2B2C2 gene cluster. As expected, the deletion mutant strains WAC9 to WAC12 carrying prpD1B1C1 and prpD2B2C2 double mutations showed no growth on propionate. Taken together, the results of the two-dimensional gel electrophoreses and the growth tests indicate that the C. glutamicum prpD2B2C2 gene cluster is essential for growth on propionate, while the prpD1B1C1 functions are not involved in propionate degradation of C. glutamicum at least under the test conditions applied.



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  		FIG. 4. Plate tests for the identification of prp genes essential for growth of C. glutamicum on propionate as sole carbon source. C. glutamicum RES167 was used as the control strain on each plate. Deletion mutant strains carrying mutations in the prpD1B1C1 region and the prpD2B2C2 region and the respective double mutant strains, corresponding to strains WAC1 to WAC12 (Table 1), were arranged on the plates by mutated prp genes. Cells (2 x 106) of each strain were spread onto a quarter of a minimal medium MM1 plate with 6 g of propionate liter-1 as the sole carbon source and incubated for 20 days at 30°C.

 

   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Two prpDBC gene clusters are present in the C. glutamicum genome. In the present study we described the cloning, sequencing, and molecular characterization of two distinct prpDBC gene clusters with high degrees of sequence similarity in C. glutamicum. Duplication of a complete prp gene cluster seems to be a unique feature of C. glutamicum, since, to our knowledge, no other bacterial genome sequenced to date is equipped with two or more chromosomal copies of prp genes. Under the growing number of genome sequences available in the databases, several prokaryotes revealed the existence of prp gene loci of different gene arrangements and sizes. This is true for both gram-negative and gram-positive eubacteria as well as for archaea. For example, the gram-positive Bacillus subtilis (GenBank accession no. NC000964) as well as Bacillus halodurans (GenBank accession no. NC002570) have a prpCDB gene cluster, whereas in Mycobacterium tuberculosis (GenBank accession no. NC000962 and NC002755) a prpDC locus is found. However, up to now no functional analyses of these prp genes have been reported, and no biochemical evidence has been published for operation of the 2-methylcitrate pathway in these gram-positive organisms.

Like C. glutamicum, some other bacteria possess three citrate synthase homologues, e.g., B. subtilis and M. tuberculosis; however, only one of these in each organism is clustered with prp genes, and their function still remains unclear. On the other hand, BLAST search results showed that some prokaryotic genomes seem to contain no prp-homologous genes (apart from their putative citrate synthase or isocitrate lyase genes involved in the citric and glyoxylic acid cycles), e.g., Corynebacterium diphtheriae (GenBank accession no. NC002935) and Mycobacterium leprae (GenBank accession no. NC002677), which are close relatives of C. glutamicum. At present it is unclear whether these organisms are able to utilize propionate at all.

Up to now, the gram-negative bacteria S. enterica, E. coli, and R. eutropha are the only other prokaryotes in which prp gene clusters are well characterized on the genetic level as well as on the biochemical level. For S. enterica, all enzymes of the propionate-degrading methylcitrate pathway—with the exception of the 2-methyl-cis-aconitate hydratase function, which is encoded by acnA or acnB—are part of the prpBCDE operon (24). In contrast to the situation for S. enterica, no prpE homologue gene encoding the propionyl-CoA synthetase was located adjacent to either or both C. glutamicum prpDBC gene loci. Thus, the activation of propionate to propionyl-CoA in C. glutamicum may be dependent on the activity of the phosphotransacetylase and the acetate kinase, encoded by the pta-ack operon, since the acetate kinase Ack was shown to be active also with propionate as substrate, and since no acetyl-CoA synthetase activity is measurable for C. glutamicum (39).

The prpC1 and prpC2 gene products have activities as both citrate and 2-methylcitrate synthases. We have presented evidence that in addition to GltA, two further proteins of C. glutamicum, PrpC1 and PrpC2, are able to mediate citrate synthase activity, although the primary cellular function of at least PrpC2 resides in the activity as 2-methylcitrate synthase induced during growth on propionate. In comparison, the citrate synthase of the psychrophilic gram-positive bacterium DS2-3R (15) showed a high amino acid sequence identity of about 50% to the C. glutamicum prpC1 and prpC2 gene products. Interestingly, in crude extracts this enzyme of DS2-3R likewise mediates citrate and 2-methylcitrate synthase activities, but the activity is independent of propionate addition to the growth substrate (14). Thus, in the absence of further genomic data, it remains unclear if the citrate synthase gene is clustered within a prp locus in DS2-3R and if propionate oxidation is dependent on 2-methylcitrate cycle activity in this organism.

Currently, all known 2-methylcitrate synthases are active on acetyl-CoA and propionyl-CoA, respectively. Interestingly, the two C. glutamicum PrpC proteins revealed different specific activities with the different CoA substrates. While PrpC1 showed an about twofold-higher level of specific activity towards acetyl-CoA than towards propionyl-CoA under standard assay conditions, PrpC2 showed nearly the reciprocal activity. Like PrpC2, the 2-methylcitrate synthases of the prokaryotes E. coli, S. enterica, and R. eutropha reveal higher specific activities with propionyl-CoA as the substrate (4, 22, 55), whereas the recently identified 2-methylcitrate synthase McsA of the fungus Aspergillus nidulans, like PrpC1 of C. glutamicum, reveals a higher specific enzyme activity towards acetyl-CoA (7). However, the C. glutamicum PrpC1 and PrpC2 proteins reveal comparable amino acid sequence similarities to the eukaryotic 2-methylcitrate synthase, with 25 and 27% identical amino acid residues, respectively.

Operation of the 2-methylcitrate pathway in C. glutamicum is dependent on the prpD2B2C2 gene cluster. We showed that C. glutamicum is able to use propionate as a sole or additional carbon and energy source, although growth revealed a relatively long lag phase. Despite its toxicity to numerous microorganisms, many fungi and bacteria are able to use propionate, even though, e.g., Saccharomyces cerevisiae is not able to use propionate as a sole carbon source (38). The catabolism of propionate seems to be useful for soil microorganisms like C. glutamicum, since propionate as the product of fermentative processes is, after acetate, the most abundant fatty acid in soil (8). Within the past years, at least six different pathways for the metabolism of propionate were postulated (17, 55). Even though it is proposed that the methylcitrate cycle may be widely distributed in bacteria for oxidative degradation of propionate (7, 22), no experimental data were published beside those for E. coli, S. enterica, R. eutropha, and, very recently, Burkholderia sacchari (5). The only further biochemical evidence for activity of this pathway in prokaryotes was the observation of 2-methylcitrate synthase activity in extracts of propionate-grown Pseudomonas aeruginosa (58).

For C. glutamicum, none of the several propionate-degrading pathways has been reported to date. Since we were able to show that the specific 2-methylcitrate synthase activity in C. glutamicum is dramatically increased during growth on propionate, that PrpC1 and PrpC2 comprised 2-methylcitrate synthase activities, and that the prpC genes are clustered with putative 2-methylisocitrate lyase (PrpB) and 2-methylcitrate dehydratase (PrpD) genes, we suggested that C. glutamicum metabolizes propionate via the 2-methylcitrate pathway. Under the test conditions applied, the crucial enzymatic functions of this pathway are encoded by the prpD2B2C2 gene cluster, since these genes were shown to be essential with propionate as the sole carbon source and since the gene products of this cluster were synthesized in strongly increased amounts in propionate-grown cells.

The function of the prpD1B1C1 gene cluster of C. glutamicum is obscure. At present, the only prp genes found to be nonessential for growth on propionate were prpE of S. enterica and prpD of R. eutropha. Since mutations within genes of the C. glutamicum prpD1B1C1 gene cluster are not linked to any phenotypic alteration, it remains debatable whether the prpD1B1C1 gene cluster is functional in C. glutamicum. Some prerequisites for successful gene expression are found for the C. glutamicum prpD1B1C1 locus, e.g., that none of the genes within this locus is interrupted by a stop codon and that a putative rho-independent termination signal as well as putative ribosome binding sites are located within the cluster. Additionally, the expression of the prpC1 gene of C. glutamicum showed clearly that the gene product is synthesized and active as a 2-methylcitrate synthase. On the other hand, propionate degradation under our test conditions is dependent on prpD2B2C2 functions, and their function cannot be assumed by prpD1B1C1. Thus, the maintenance of the prpD1B1C1 gene cluster must be based on some selective pressure, since otherwise it would have been lost.

Recent findings revealed evidence for the existence of at least two pathways involved in propionic acid oxidation in B. sacchari during production of poly-3-hydroxybutyrate-co-3-hydroxyvalerate (44). It was shown very recently that in this organism the 2-methylcitrate cycle operates at relatively low propionate concentrations (0.2 to 0.4 g of propionate liter-1), while a second, hitherto unidentified pathway is more important at higher propionate concentrations (5). Thus, the expression of the C. glutamicum prpD1B1C1 gene clusters may also be dependent on the ambient propionate concentration.

Another possible explanation for the function of the prpD1B1C1 locus might be the involvement in a hitherto unknown metabolic pathway. A Legionella pneumophila prpD transposon insertion mutant was shown to be unaffected during growth in vitro but revealed reduced cytopathogenicity to macrophage-like cell lines as well as impaired intracellular survival and replication within macrophages and amoebae (47). The same authors suggest a biosynthetic role for the prp locus of L. pneumophila. Thus, even though the prpD2B2C2 gene cluster was shown to be necessary for propionate degradation via the 2-methylcitrate cycle for C. glutamicum, the role of the prpD1B1C1 genes remains to be analyzed.


   ACKNOWLEDGMENTS
 
We thank Bernhard J. Eikmanns for kindly providing the plasmid pUC18-gltA, the Degussa AG for providing nucleotide sequence data and financial support, and Wolfgang Buckel for critical reading of the manuscript.


   FOOTNOTES
 
* Corresponding author. Mailing address: Lehrstuhl für Genetik, Fakultät für Biologie, Universität Bielefeld, Postfach 100131, 33501 Bielefeld, Germany. Phone: 49-521-106-4825. Fax: 49-521-106-5626. E-mail: Joern.Kalinowski{at}Genetik.Uni-Bielefeld.DE. Back


   REFERENCES
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Altschul, S. F., T. L. Madden, A. A. Schaeffer, J. Zhang, Z. Zhang, W. Miller, and D. J. Lipman. 1997. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res. 25:3389-3402.[Abstract/Free Full Text]
  2. Amador, E., J. M. Castro, A. Correira, and J. F. Martin. 1999. Structure and organization of the rrnD operon of Brevibacterium lactofermentum: analysis of the 16S rRNA gene. Microbiology 145:915-924.[Abstract]
  3. Amann, E., B. Ochs, and K. J. Abel. 1988. Tightly regulated tac promoter vectors useful for the expression of unfused and fused proteins in Escherichia coli. Gene 69:301-315.[CrossRef][Medline]
  4. Brämer, C. O., and A. Steinbüchel. 2001. The methylcitric acid pathway in Ralstonia eutropha: new genes identified involved in propionate metabolism. Microbiology 147:2203-2214.[Abstract/Free Full Text]
  5. Brämer, C. O., L. F. Silva, J. G. C. Gomez, H. Priefert, and A. Steinbüchel. 2002. Identification of the 2-methylcitrate pathway involved in the catabolism of propionate in the polyhydroxyalkanoate-producing strain Burkholderia sacchari IPT101T and analysis of a mutant accumulating a copolyester with higher 3-hydroxyvalerate content. Appl. Environ. Microbiol. 68:271-279.[Abstract/Free Full Text]
  6. Brock, M., D. Darley, S. Textor, and W. Buckel. 2001. 2-Methylisocitrate lyase from the bacterium Escherichia coli and the filamentous fungus Aspergillus nidulans. Characterization and comparison of both enzymes. Eur. J. Biochem. 268:3577-3586.[Medline]
  7. Brock, M., R. Fischer, D. Linder, and W. Buckel. 2000. Methylcitrate synthase from Aspergillus nidulans: implications for propionate as an antifungal agent. Mol. Microbiol. 35:961-973.[CrossRef][Medline]
  8. Buckel, W. 1999. Anaerobic energy metabolism, p. 278-326. In J. W. Lengler, G. Drews, H. G. Schlegel (ed.), Biology of the procaryotes. Thieme, Stuttgart, Germany.
  9. Cocaign-Bousquet, M., A. Guyonvarch, and N. D. Lindley. 1996. Growth rate-dependent modulation of carbon flux through the central metabolism and the kinetic consequences for glucose-limited chemostat cultures of Corynebacterium glutamicum. Appl. Environ. Microbiol. 62:429-436.[Abstract]
  10. Degussa, A. G. November 1994. European patent 0 375 889.
  11. Dominguez, H., C. Nezondet, N. D. Lindley, and M. Cocaign. 1993. Modified carbon flux during oxygen limited growth of Corynebacterium glutamicum and the consequences for amino acid overproduction. Biotechnol. Lett. 15:449-454.[CrossRef]
  12. Eikmanns, B. J., N. Thum-Schmitz, L. Eggeling, K. U. Lüdtke, and H. Sahm. 1994. Nucleotide sequence, expression and transcriptional analysis of the Corynebacterium glutamicum gltA gene encoding citrate synthase. Microbiology 140:1817-1828.[Abstract]
  13. Gaasterland, T., and C. W. Sensen. 1996. Fully automated genome analysis that reflects user needs and preferences. A detailed introduction to the MAGPIE system architecture. Biochimie 78:302-310.[Medline]
  14. Gerike, U., D. W. Hough, N. J. Russel, M. L. Dyall-Smith, and M. J. Danson. 1998. Citrate synthase and 2-methylcitrate synthase: structural, functional and evolutionary relationships. Microbiology 144:929-935.[Abstract]
  15. Gerike, U., M. J. Danson, N. J. Russel, and D. W. Hough. 1997. Sequencing and expression of the gene encoding a cold-active citrate synthase from an Antarctic bacterium, strain DS2-3R. Eur. J. Biochem. 248:49-57.[Medline]
  16. Grant, S. G., J. Jessee, F. R. Bloom, and D. Hanahan. 1990. Differential plasmid rescue from transgenic mouse DNAs into Escherichia coli methylation-restriction mutants. Proc. Natl. Acad. Sci. USA 87:4645-4649.[Abstract/Free Full Text]
  17. Halarnkar, P. P., and G. J. Blomquist. 1989. Comparative aspects of propionate metabolism. Comp. Biochem. Physiol. 92:227-231.[CrossRef]
  18. Haynes, J. A., and M. L. Britz. 1989. Electrotransformation of Brevibacterium lactofermentum and Corynebacterium glutamicum: growth in Tween 80 increases transformation frequencies. FEMS Microbiol. Lett. 61:329-334.[CrossRef]
  19. Hermann, T., W. Pfefferle, C. Baumann, E. Busker, S. Schaffer, M. Bott, H. Sahm, N. Dusch, J. Kalinowski, A. Pühler, A. K. Bendt, R. Krämer, and A. Burkovski. 2001. Proteome analysis of Corynebacterium glutamicum. Electrophoresis 22:1712-1723.[CrossRef][Medline]
  20. Hodgson, J. 1998. LION and Degussa apply genomics to fermentation. Nat. Biotechnol. 16:715.[CrossRef][Medline]
  21. Horswill, A. R., and J. C. Escalante-Semerena. 1997. Propionate catabolism in Salmonella typhimurium LT2: two divergently transcribed units comprise the prp locus at 8.5 centisomes, prpR encodes a member of the sigma-54 family of activators, and the prpBCDE genes constitute an operon. J. Bacteriol. 179:928-940.[Abstract/Free Full Text]
  22. Horswill, A. R., and J. C. Escalante-Semerena. 1999. Salmonella typhimurium LT2 catabolizes propionate via the 2-methylcitric acid cycle. J. Bacteriol. 181:5615-5623.[Abstract/Free Full Text]
  23. Horswill, A. R., and J. C. Escalante-Semerena. 1999. The prpE gene of Salmonella typhimurium LT2 encodes propionyl-CoA synthetase. Microbiology 145:1381-1388.[Abstract]
  24. Horswill, A. R., and J. C. Escalante-Semerena. 2001. In vitro conversion of propionate by Salmonella enterica enzymes: 2-methylcitrate dehydratase (PrpD) and aconitase enzymes catalyze the conversion of 2-methylcitrate to 2-methylisocitrate. Biochemistry 40:4703-4713.[CrossRef][Medline]
  25. Horton, R. M. 1995. PCR mediated recombination and mutagenesis. SOEing together tailor-made genes. Mol. Biotechnol. 3:93-99.[Medline]
  26. Jones, D., and M. D. Collins. 1986. Section 15. Irregular, nonsporing Gram-positive rods, p. 1261-1434. In P. H. A. Sneath, N. S. Mair, M. E. Sharpe, and G. H. Holt (ed.), Bergey's manual of systematic bacteriology, vol. 2. Williams & Wilkins, Baltimore, Md.
  27. Katsumata, R., A. Ozaki, T. Oka, and A. Furuya. 1984. Protoplast transformation of glutamate-producing bacteria with plasmid DNA. J. Bacteriol. 159:306-311.[Abstract/Free Full Text]
  28. Keilhauer, C., L. Eggeling, and H. Sahm. 1993. Isoleucin synthesis in Corynebacterium glutamicum: molecular analysis of the ilvB-ilvN-ilvC operon. J. Bacteriol. 175:5595-5603.[Abstract/Free Full Text]
  29. Leuchtenberger, W. 1996. Amino acids: technical production and use, p. 465-502. In H. J. Rehm, G. Reed, A. Pühler, and P. Stadler (ed.), Bio/Technology. VCH, Weinheim, Germany.
  30. London, R. E., D. L. Allen, S. A. Gabel, and E. F. DeRose. 1999. Carbon-13 nuclear magnetic resonance study of metabolism of propionate by Escherichia coli. J. Bacteriol. 181:3562-3570.[Abstract/Free Full Text]
  31. Luttik, M. A. H., P. Kötter, F. A. Salomons, I. J. van der Klei, J. P. van Dijken, and J. T. Pronk. 2000. The Saccharomyces cerevisiae ICL2 gene encodes a mitochondrial 2-methylisocitrate lyase involved in propionyl-coenzyme A metabolism. J. Bacteriol. 182:7007-7013.[Abstract/Free Full Text]
  32. Miyakoshi, S., H. Uchiyama, T. Someya, T. Satoh, and T. Tabuchi. 1987. Distribution of the methylcitric acid cycle and ß-oxidation pathway for propionate catabolism in fungi. Agric. Biol. Chem. 51:2381-2387.
  33. Norrander, J., T. Kempe, and J. Messing. 1983. Construction of improved M13 vectors using oligonucleotide-directed mutagenesis. Gene 26:101-106.[CrossRef][Medline]
  34. O'Farrell, P. H. 1975. High-resolution two-dimensional electrophoresis of proteins. J. Biol. Chem. 250:4007-4021.[Abstract/Free Full Text]
  35. Oppenheim, D. S., and C. Yanofski. 1980. Translational coupling during expression of the tryptophan operon of Escherichia coli. Genetics 95:785-795.[Abstract/Free Full Text]
  36. Palacios, S., and J. C. Escalante-Semerena. 2000. prpR, ntrA, and ihf functions are required for expression of the prpBCDE operon, encoding enzymes that catabolize propionate in Salmonella enterica serovar Typhimurium LT2. J. Bacteriol. 182:905-910.[Abstract/Free Full Text]
  37. Perkins, D. N., D. J. Pappin, D. M. Creasy, and J. S. Cottrell. 1999. Probability-based protein identification by searching sequence databases using mass spectrometry data. Electrophoresis 20:3551-3567.[CrossRef][Medline]
  38. Pronk, J. T., A. van der Linden-Beuman, C. Verduyn, W. A. Scheffers, and J. P. van Dijken. 1994. Propionate metabolism in Saccharomyces cerevisiae: implications for the metabolon hypothesis. Microbiology 140:717-722.[Abstract]
  39. Reinscheid, D. J., S. Schnicke, D. Rittmann, U. Zahnow, H. Sahm, and B. J. Eikmanns. 1999. Cloning, sequence analysis, expression and inactivation of the Corynebacterium glutamicum pta-ack operon encoding phosphotransacetylase and acetate kinase. Microbiology 145:503-513.[Abstract]
  40. Sahm, H., L. Eggeling, and A. A. de Graaf. 2000. Pathway analysis and metabolic engineering in Corynebacterium glutamicum. Biol. Chem. 381:899-910.[CrossRef][Medline]
  41. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
  42. Schäfer, A., A. Tauch, W. Jäger, J. Kalinowski, G. Thierbach, and A. Pühler. 1994. Small mobilizable multi-purpose cloning vectors derived from the Escherichia coli plasmid pK18 and pK19: selection of defined deletions in the chromosome of Corynebacterium glutamicum. Gene 145:69-73.[CrossRef][Medline]
  43. Shiio, I., H. Ozaki, and K. Ujigawa-Takeda. 1982. Production of aspartic acid and lysine by citrate synthase mutants of Brevibacterium flavum. Agric. Biol. Chem. 46:101-107.
  44. Silva, L. F., J. G. C. Gomez, M. S. Olivieira, and B. B. Torres. 2000. Propionic acid metabolism and poly-3-hydroxybutyrate-co-3-hydroxyvalerate (P3HB-co-3HV) production by Burkholderia sp. J. Biotechnol. 76:165-174.[CrossRef][Medline]
  45. Srere, P. A. 1969. Citrate synthase. Methods Enzymol. 13:3-11.
  46. Staden, R. 1996. The Staden sequence analysis package. Mol. Biotechnol. 5:233-241.[Medline]
  47. Stone, B. J., A. Brier, and Y. A. Kwaik. 1999. The Legionella pneumophila prp locus; required during infection of macrophages and amoebae. Microb. Pathog. 27:369-376.[CrossRef][Medline]
  48. Suggs, S. V., T. Hirose, T. Miyake, E. H. Kawashima, M. J. Johnson, K. Itakura, and R. B. Wallace. 1981. Use of synthetic oligodeoxyribonucleotides for the isolation of specific cloned DNA sequences, p. 683-693. In D. D. Brown and C. F. Fox (ed.), Developmental biology using purified genes. Academic Press, Inc., New York, N.Y.
  49. Tabuchi, T., and H. Uchiyama. 1975. Methylcitrate condensing and methylisocitrate cleaving enzymes; evidence for the pathway of oxidation of propionyl-CoA to pyruvate via C7-tricarboxylic acids. Agric. Biol. Chem. 39:2035-2042.
  50. Tabuchi, T., and S. Hara. 1974. Production of 2-methylcitric acid from n-paraffins by mutants of Candida lipolytica. Agric. Biol. Chem. 38:1105-1106.
  51. Tabuchi, T., H. Aoki, H. Uchiyama, and T. Nakahara. 1981. 2-Methylcitric dehydratase, a new enzyme functioning at the methylcitric acid cycle of propionate metabolism. Agric. Biol. Chem. 45:2823-2829.
  52. Tauch, A., F. Kassing, J. Kalinowski, and A. Pühler. 1995. The Corynebacterium xerosis composite transposon Tn 5432 consists of two identical insertion sequences, designated IS 1249, flanking the erythromycin resistance gene ermCX. Plasmid 34:119-131.[CrossRef][Medline]
  53. Tauch, A., I. Homann, S. Mormann, S. Rüberg, A. Billault, B. Bathe, S. Brand, O. Brockmann-Gretza, C. Rückert, N. Schischka, C. Wrenger, J. Hoheisel, B. Möckel, K. Huthmacher, W. Pfefferle, A. Pühler, and J. Kalinowski. 2002. Strategy to sequence the genome of Corynebacterium glutamicum ATCC 13032: use of a cosmid and a bacterial artificial chromosome library. J. Biotechnol. 95:25-38.[CrossRef][Medline]
  54. Tauch, A., O. Kirchner, L. Wehmeier, J. Kalinowski, and A. Pühler. 1994. Corynebacterium glutamicum DNA is subjected to methylation-restriction in Escherichia coli. FEMS Microbiol. Lett. 123:343-348.[CrossRef][Medline]
  55. Textor, S., V. F. Wendisch, A. A. De Graaf, U. Müller, M. I. Linder, D. Linder, and W. Buckel. 1997. Propionate oxidation in Escherichia coli: evidence for operation of a methylcitrate cycle in bacteria. Arch. Microbiol. 168:428-436.[CrossRef][Medline]
  56. Thompson, J. D., D. G. Higgins, and T. J. Gibson. 1994. CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res. 22:4673-4680.[Abstract/Free Full Text]
  57. Thomson, J. M., and W. A. Parrott. 1998. pMECA: a cloning plasmid with 44 unique restriction sites that allows selection of recombinants based on colony size. BioTechniques 24:922-924.[Medline]
  58. Watson, D., D. L. Lindel, and R. Fall. 1983. Pseudomonas aeruginosa contains an inducible methylcitrate synthase. Curr. Microbiol. 8:17-21.
  59. Wendisch, V. F., A. A. de Graaf, H. Sahm, and B. J. Eikmanns. 2000. Quantitative determination of metabolic fluxes during co-utilization of two carbon sources: comparative analyses with Corynebacterium glutamicum during growth on acetate and/or glucose. J. Bacteriol. 182:3088-3096.[Abstract/Free Full Text]
  60. Wohlleben, W., G. Muth, and J. Kalinowski. 1993. Genetic engineering of Gram positive bacteria, p. 457-505. In H. J. Rehm, G. Reed, A. Pühler, and P. Stadler (ed.), Bio/Technology. VCH, Weinheim, Germany.
  61. Zuker, M., D. H. Mathews, and D. H. Turner. 1999. Algorithms and thermodynamics for RNA secondary structure prediction: a practical guide in RNA biochemistry and biotechnology, p. 11-43. J. Barciszewski and B. F. C. Clark (ed.), NATO ASI series. Kluwer Academic Publishers, New York, N.Y.

Journal of Bacteriology, May 2002, p. 2728-2739, Vol. 184, No. 10
0021-9193/02/$04.00+0     DOI: 10.1128/JB.184.10.2728-2739.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.



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