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Journal of Bacteriology, November 2006, p. 7668-7676, Vol. 188, No. 21
0021-9193/06/$08.00+0     doi:10.1128/JB.01009-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

Knockout and Overexpression of Pyrroloquinoline Quinone Biosynthetic Genes in Gluconobacter oxydans 621H{triangledown}

Tina Hölscher* and Helmut Görisch

Fachgebiet Technische Biochemie, Institut für Biotechnologie, Technische Universität Berlin, D-13353 Berlin, Germany

Received 10 July 2006/ Accepted 17 August 2006


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ABSTRACT
 
In Gluconobacter oxydans, pyrroloquinoline quinone (PQQ) serves as the cofactor for various membrane-bound dehydrogenases that oxidize sugars and alcohols in the periplasm. Proteins for the biosynthesis of PQQ are encoded by the pqqABCDE gene cluster. Our reverse transcription-PCR and promoter analysis data indicated that the pqqA promoter represents the only promoter within the pqqABCDE cluster of G. oxydans 621H. PQQ overproduction in G. oxydans was achieved by transformation with the plasmid-carried pqqA gene or the complete pqqABCDE cluster. A G. oxydans mutant unable to produce PQQ was obtained by site-directed disruption of the pqqA gene. In contrast to the wild-type strain, the pqqA mutant did not grow with D-mannitol, D-glucose, or glycerol as the sole energy source, showing that in G. oxydans 621H, PQQ is essential for growth with these substrates. Growth of the pqqA mutant, however, was found with D-gluconate as the energy source. The growth behavior of the pqqA mutant correlated with the presence or absence of the respective PQQ-dependent membrane-bound dehydrogenase activities, demonstrating the vital role of these enzymes in G. oxydans metabolism. A different PQQ-deficient mutant was generated by Tn5 transposon mutagenesis. This mutant showed a defect in a gene with high homology to the Escherichia coli tldD gene, which encodes a peptidase. Our results indicate that the tldD gene in G. oxydans 621H is involved in PQQ biosynthesis, possibly with a similar function to that of the pqqF genes found in other PQQ-synthesizing bacteria.


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INTRODUCTION
 
The acetic acid bacterium Gluconobacter oxydans is characterized by its ability to incompletely oxidize various sugars and alcohols by using membrane-associated dehydrogenases that contain pyrroloquinoline quinone (PQQ) as a cofactor (33). Examples are the quinoprotein glucose dehydrogenase (3) and the quinoprotein glycerol dehydrogenase, which also oxidizes D-gluconate, D-mannitol, D-sorbitol, and other polyols (34). The oxidation reactions take place in the periplasmic space and are coupled to the respiratory chain (19, 33). Several oxidation reactions carried out by G. oxydans quinoproteins are of industrial importance, e.g., the conversion of D-gluconate to 5-D-ketogluconate, a precursor for the production of L-tartaric acid, and the formation of L-sorbose, an intermediate in the synthesis of vitamin C (reviewed in reference 9). PQQ has invoked considerable attention due to its positive physiological effects in mammals (47). A possible role of PQQ as a vitamin in mammals has been suggested but is controversially debated (13, 28, 40).

Genes involved in PQQ synthesis have been characterized for several bacteria, including Klebsiella pneumoniae, Acinetobacter calcoaceticus, Methylobacterium extorquens AM1, and Pseudomonas sp. (reviewed in reference 19). In Klebsiella pneumoniae, the PQQ biosynthetic genes are clustered in the pqqABCDEF operon (35). In Pseudomonas aeruginosa, the pqqABCDE operon is separated from the pqqF operon (18, 48). Methylobacterium extorquens AM1 contains a pqqABC/DE operon in which the pqqC and pqqD genes are fused, while the pqqFG genes form an operon with three other genes (36, 46, 54). In Acinetobacter calcoaceticus, a pqqABCDE cluster but no pqqF gene is known (20). The details of PQQ biosynthesis have not yet been resolved; however, some of the involved proteins have been functionally characterized. The pqqA genes from different species encode small peptides of 23 to 29 amino acids with conserved glutamate and tyrosine residues. The PQQ backbone is constructed from glutamate and tyrosine (26, 50), and most probably these amino acids are derived from the PqqA peptide (21, 50). The PqqB protein might be involved in transport of PQQ into the periplasm (52). PqqC catalyzes the final step in PQQ formation (31). The functions of PqqD and PqqE are still unknown. The PqqF and PqqG proteins share similarity with a family of divalent cation-containing endopeptidases that cleave small peptides (35, 46). Thus, it has been speculated that PqqF and PqqG are involved in processing the peptide precursor of PQQ. While PqqF of Klebsiella pneumoniae is most closely related to the Escherichia coli peptidase pitrilysin, the PqqF and PqqG proteins of Methylobacterium extorquens show some similarity to the two subunits of mitochondrial processing peptidases (35, 46). The operon nature of the pqqABCDE(F) cluster was initially inferred from sequence analysis. In Methylobacterium extorquens, cotranscription of pqqAB (39) as well as cotranscription of each pair of genes in the pqqABC/DE cluster was shown, supporting the hypothesis that expression of the pqq cluster occurs from a single promoter upstream of pqqA (54). However, the pqqA gene, which encodes the putative PQQ precursor, is transcribed at significantly higher levels than the other pqq genes within the operon (39, 52). This was ascribed to a potential termination signal located downstream of pqqA that reduces transcription of the following genes.

Little information is available on the PQQ biosynthetic genes in G. oxydans. Gupta and coworkers (22) used Tn5 transposon mutagenesis to generate a mutant of Gluconobacter oxydans IFO 3293 with a defect in quinoprotein glucose dehydrogenase activity. Later it was shown that the lack of glucose dehydrogenase activity was due to a Tn5 insertion in the pqqE gene that prevented synthesis of the PQQ cofactor (12). This information led to the identification of the pqqABCDE cluster in G. oxydans ATCC 9937 (12), which showed the same arrangement of genes as the PQQ biosynthetic operons in other species. It was suggested, based on sequence analysis, that the pqqA gene carries its own promoter while the pqqBCDE genes are controlled by an additional weak promoter present in the large pqqA-pqqB intergenic region. However, there is no experimental evidence so far for the presence of additional promoters within the pqqABCDE clusters of other species. The recent sequencing of the whole genome of Gluconobacter oxydans 621H (38), the strain investigated in this study, revealed that this strain carries a pqqABCDE operon (Fig. 1A) which shares high sequence similarity with the pqqABCDE operon characterized for G. oxydans ATCC 9937. Similar to the case for Acinetobacter calcoaceticus, an equivalent of the pqqF gene has not been recognized in both G. oxydans strains so far. The aim of our study was to investigate the genes involved in PQQ biosynthesis and the relevance of the cofactor PQQ for growth and oxidizing activities of G. oxydans 621H.


Figure 1
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  		FIG. 1. Genetic organization of the pqqABCDE cluster (A) and the putative tldD-containing operon (B) in G. oxydans 621H. P, putative promoter region. Numbers refer to gene loci of the G. oxydans 621H genome sequence (accession number CP000009).


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MATERIALS AND METHODS
 
Bacterial strains, plasmids, and culture conditions. The relevant characteristics and references for the strains and plasmids used in this work are listed in Table 1. E. coli strains were grown in Luria-Bertani medium. Gluconobacter oxydans ATCC 621H was cultivated routinely in complex medium containing 0.5% yeast extract and 0.3% tryptone, pH 6.0, supplemented with 5% D-mannitol (mannitol medium). When indicated, D-mannitol was replaced by other sources of carbon and energy (D-glucose, D-gluconate, or glycerol). To study PQQ production, cultures were grown in a minimal medium containing salts, vitamins, and 0.2 g/liter L-glutamine (37) supplemented with 5% D-mannitol or with 2% D-mannitol and 2% D-gluconate. When appropriate, a medium based on the minimal medium, but supplemented with small amounts of yeast extract (0.05%) and tryptone (0.03%), was used. G. oxydans was cultivated aerobically at 150 rpm and 30°C. Growth was recorded by optical density measurements at 620 nm. For selection of potential G. oxydans PQQ-deficient mutants, colonies were transferred to glucose-calcium carbonate agar (10% D-glucose, 5% yeast extract, 0.5 g/liter MgSO4, 10 g/liter CaCO3) (22), and the absence of clearing zones around the colonies was recorded. Antibiotics were used at the following concentrations: ampicillin, 100 µg/ml; cefoxitin, 50 µg/ml; gentamicin, 10 µg/ml (E. coli) or 50 µg/ml (G. oxydans); kanamycin, 50 µg/ml; and tetracycline, 20 µg/ml.


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  		TABLE 1. Strains and plasmids used in this study

General genetic techniques. DNA work was performed according to standard protocols (5, 43). For PCRs, genomic DNA isolated from G. oxydans ATCC 621H was used as a template. Pfu DNA polymerase (Promega) was used for PCR amplification of inserts for cloning, and Taq DNA polymerase (Rapidozym) was used for PCR amplification in test reactions (e.g., colony PCR) and reverse transcriptase PCR (RT-PCR). The primers used in this study are listed in Table 2. For primer design, the whole genome sequence of G. oxydans 621H (38; http://www.ncbi.nlm.nih.gov [accession number CP000009]) was used.


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  		TABLE 2. Primers used in this study

RNA preparation and RT-PCR. For RNA isolation, G. oxydans was grown overnight in mannitol medium. Total RNA was isolated from 1 ml of culture, using a QIAGEN RNeasy mini kit according to the manufacturer's instructions. To remove residual DNA, RNA was treated with DNase I for 3 h at 37°C and subsequently cleaned up using the RNeasy kit reagents. Reverse transcription was carried out in 20-µl reaction mixtures containing 5 µg RNA, 1 µM reverse primer BR2, 0.5 M betaine, a 0.5 mM concentration of each deoxynucleoside triphosphate, 20 U Ribolock RNase inhibitor, 10 mM dithiothreitol, and 200 U SuperScript III RT (Invitrogen) in first-strand buffer (Invitrogen). After incubation of the prehybridization mix (RNA, primer, and betaine) at 65°C for 5 min, all other components were added. Reverse transcription was carried out at 48°C for 30 min, 50°C for 20 min, 55°C for 10 min, and 70°C for 15 min. Subsequently, the RNA was digested with 5 U RNase H (New England Biolabs) at 37°C for 20 min. As no-RT controls, reaction mixtures were set up with all ingredients except the reverse transcriptase. The resulting cDNA was used directly for RT-PCR.

Construction of expression vectors. A 3.7-kb DNA fragment of G. oxydans 621H bearing the pqqABCDE operon (38) (Fig. 1A) and parts of the upstream and downstream open reading frames was amplified by PCR with primers PQQ-F-eco and PQQ-R-xbaI and cloned between the EcoRI-XbaI sites of pUC18, resulting in plasmid pTB9019. To generate plasmid pTB9028, the pqqABCDE operon was excised from pTB9019 and cloned between the EcoRI-HindIII sites of plasmid pBBR1MCS-5. A 520-bp fragment containing the pqqA gene, including parts of the upstream gene, was amplified with primers PQQ-F-eco and pqqA-HindR and cloned between the EcoRI-HindIII sites of pBBR1MCS-5, resulting in plasmid pTB9031. A 4.1-kb fragment containing the putative tldD-containing operon of G. oxydans 621H (Fig. 1B; see Results for details) was amplified using primers hpF-xho and tldR-xba and cloned between the XhoI-XbaI sites of pBBR1MCS-5, resulting in plasmid pTB9045. For expression of the tldD gene alone, the promoter of the elongation factor (tufB promoter) (4, 41) was used. The tufB promoter was amplified from G. oxydans 621H, using the primers tuG-SalF and tuG-HindR, and cloned into the SalI-HindIII sites of pBBR1MCS-5. The tldD gene (1.8 kb) was amplified using primers tldF-hind and tldR-xba, and the fragment was cloned between the HindIII-XbaI sites downstream of the tufB promoter, resulting in plasmid pTB9046. To generate plasmid pTB9055, the pqqABCDE operon was cloned into the KpnI-XhoI sites of pTB9046, upstream of the tufB promoter and the tldD gene.

Construction of promoter-probe vectors. The region upstream of pqqA (300 bp) and the pqqA-pqqB intergenic region (180 bp) containing the putative pqqA and pqqBCDE promoters (12) were amplified by PCR, using the primer pairs PpqqAF/PpqqArev1 and PpqqBF/PpqqBrev1, respectively. The amplicons were cloned between the EcoRI-HindIII sites of the promoter-probe vector pCM130 (32), resulting in plasmids pTB9026 (pqqBCDE promoter region) and pTB9027 (pqqA promoter region).

Construction of pqqA and tldD knockout vectors. To generate a gene replacement vector for inactivation of the pqqA gene, the strategy of Derbise et al. (10) was adapted. A DNA fragment containing the kanamycin resistance gene of transposon Tn5 inserted between the pqqA promoter region and the pqqB gene, resulting in almost complete deletion of pqqA, was generated by PCR as follows. A 500-bp fragment containing the region upstream of pqqA and a 700-bp fragment containing the region downstream of pqqA were amplified by PCR, using primer pairs Up-F/Up-R-kan and Down-F-kan/Down-R, respectively. The flanking regions were fused to the ends of the kanamycin resistance cassette of transposon Tn5 by two consecutive PCRs with primers Up-F and Down-R, with the products of the first PCR used as templates in the second PCR. The second PCR resulted in multiple bands. The prominent, 2.2-kb band was excised from the gel, purified, and cloned into the EcoRI-HindIII sites of pEX18Ap, resulting in plasmid pTB9038.

To generate a gene replacement vector for inactivation of the tldD gene, the tldD gene of G. oxydans 621H was cloned between the HindIII-XbaI sites of pKmob18GII, resulting in plasmid pTB9047. The tldD gene was interrupted by insertion of the gentamicin resistance cassette of vector pBBR1MCS-5 into the NotI-AatII sites of pTB9047, resulting in plasmid pTB9048.

Conjugational plasmid transfer into G. oxydans. Expression vectors and knockout vectors were transferred into G. oxydans by triparental mating, using E. coli DH5{alpha} bearing the respective vector as the donor and E. coli HB101 bearing plasmid pRK2013 as the helper strain. The three strains were grown to late exponential phase, pelleted, resuspended in mannitol medium, and mixed at a 1:1:1 ratio. The mixture was plated on mannitol medium agar and incubated overnight at 30°C. The grown cell patches were scraped from the plates and streaked on selective mannitol medium agar containing cefoxitin and the appropriate selective antibiotic (kanamycin or gentamicin). Plates were incubated for 2 to 4 days until kanamycin- or gentamicin-resistant colonies appeared.

Tn5 mutagenesis. Transposon mutagenesis in G. oxydans 621H was carried out by conjugational transfer of vector pSUP1021 bearing transposon Tn5 (44) into G. oxydans. The E. coli S17-1 donor strain containing pSUP1021 and the recipient G. oxydans strain were grown to exponential phase, pelleted, and resuspended in mannitol medium. A 1:2 mixture of donor and recipient was plated on mannitol medium agar and incubated overnight at 30°C. The grown cell patches were scraped from the plates, resuspended in mannitol medium to a cell number of 108 to 109 per ml, and plated on selective mannitol medium agar containing cefoxitin and kanamycin. Plates were incubated for 2 to 4 days until kanamycin-resistant colonies appeared.

Inverse PCR. To locate the transposon Tn5 insertion site in the TH2 mutant, the strategy described by Huang et al. (27), based on inverse PCR, was carried out. As a modification, primer BR (27) was used in combination with primer BRf1Bam (Table 2).

Preparation of membrane fractions. G. oxydans strains were grown to an optical density of 0.9 in complex medium containing 2% D-gluconate and 2% D-mannitol. Cells were harvested by centrifugation at 5,000 x g for 10 min, washed once with 10 mM Tris-HCl buffer, pH 7.0, and resuspended in the same buffer. Cells were broken by three cycles of ultrasonication using a Branson sonifier 250 (intensity 4, 20% duty cycle, 5 min). After centrifugation at 10,000 x g for 10 min to remove cell debris, the supernatant was centrifuged at 100,000 x g for 60 min. The pellet was washed once, resuspended in the same buffer, and regarded as the membrane fraction.

Catechol dioxygenase assay. Cells were collected from 2 ml of culture medium by centrifugation, and the pellets were washed once in 10 mM potassium phosphate, pH 7.5, and then resuspended in 1 ml assay buffer containing 100 mM potassium phosphate, pH 7.5, and 10% acetone. Cell suspensions were diluted in assay buffer or assayed directly. The reaction was started by the addition of 10 µl 0.1 M catechol. Catechol dioxygenase activity was determined spectrophotometrically at 30°C as described previously (42, 55). Before determination of protein concentrations, cell suspensions were incubated for 20 min at 80°C.

Detection of PQQ. The presence of PQQ in culture supernatants was determined with crude membranes from E. coli containing apo-glucose dehydrogenase (16). For detection of PQQ in cells, cells were collected from 2 ml of the culture medium by centrifugation and resuspended in 200 µl MilliQ water. After incubation for 20 min at 80°C and centrifugation, the supernatant was used for PQQ detection.

Dehydrogenase assay. Dehydrogenase activities were determined photometrically at 25°C in a dye-linked system containing 2,6-dichlorophenol indophenol (DCPIP) and phenazine methosulfate. The reaction mixture contained enzyme solution, McIlvaine buffer, 33 mM substrate, 0.67 mM phenazine methosulfate, 0.1 mM DCPIP, and 4 mM sodium azide. One unit of dehydrogenase activity was defined as the reduction of 1 µmol DCPIP per min, corresponding to the oxidation of 1 µmol substrate per min. Mannitol, glycerol, and gluconate dehydrogenase activities were measured at 520 nm and pH 5.0, using a millimolar extinction coefficient of DCPIP of 10.5. Glucose dehydrogenase activity was measured at 600 nm and pH 6.0, using a millimolar extinction coefficient of DCPIP of 17.2. For reconstitution of quinoprotein apoenzymes, samples were incubated with 16.5 µM PQQ and 10 mM MgSO4 or CaCl2 for 10 min at 25°C.

Protein determination. Protein concentrations were determined as described by Bradford (7), using bovine serum albumin as the standard.

Sequence analysis. The genome sequence of G. oxydans 621H was obtained from the NCBI web site (http://www.ncbi.nlm.nih.gov [accession number CP000009]). Sequences were compared to other published sequences by using the National Center for Biotechnology Information BLASTP search tool (http://www.ncbi.nlm.nih.gov/BLAST/). Amino acid sequences were aligned with the ClustalW program located at the European Bioinformatics Institute website (http://www.ebi.ac.uk/clustalw/). Analysis of putative promoter regions was carried out with the Neural Network Promoter Prediction tool (http://www.fruitfly.org/seq_tools/promoter.html). The molecular weight of TldD was calculated using the ExPASy Compute pI/Mw tool (http://www.expasy.org/tools/pi_tool.html).


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RESULTS
 
PQQ production by wild-type G. oxydans. G. oxydans 621H excreted significant amounts of PQQ into the medium. For determination of the amount of PQQ, the strain was grown in minimal medium containing 5% D-mannitol. PQQ measured in cells resuspended in the original culture volume amounted to only about 20% of the PQQ concentration found in the culture supernatant. The PQQ concentration in the supernatant increased during growth, from about 10 ng/ml at an optical density of 0.2 to about 70 ng/ml at an optical density of 0.7.

Transcription of pqqA and pqqBCDE genes. To study the transcriptional regulation of the pqq genes in G. oxydans 621H, the promoter-probe vectors pTB9026 (putative pqqBCDE promoter) and pTB9027 (putative pqqA promoter) were transferred into the wild-type strain. After growth to an optical density of 0.4 to 0.6 in complex medium containing 5% D-mannitol or 2% D-gluconate, the catechol dioxygenase (xylE) activity was determined. With both growth substrates, the activity of the pqqBCDE promoter (<1 nmol mg–1 min–1) was below the activity found for the empty pCM130 vector (4.5 nmol mg–1 min–1). In contrast, the pqqA promoter (pTB9027) showed at least 3.5-fold higher activity (16 nmol mg–1 min–1) than the empty vector.

Reverse transcription of G. oxydans RNA was carried out using a primer targeting the pqqB open reading frame (BR2) (Fig. 2A). RT-PCR showed that the pqqA gene and the pqqB gene were cotranscribed (Fig. 2B). Using BR2-generated cDNA as the template, amplicons of 160 bp and 260 bp were generated with primer pairs AF2/BR2 and AF3/BR2, respectively.


Figure 2
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  		FIG. 2. RT-PCR of pqqAB. (A) Organization of the regulatory region of pqq genes in G. oxydans 621H and primers used for RT-PCR. PpqqA, putative pqqA promoter; PpqqB, putative pqqBCDE promoter; AF2 and AF3, forward primers used for RT-PCR; BR2, reverse transcription primer and reverse primer used for RT-PCR. (B) RT-PCR results. For RT-PCR, primers AF3/BR2 (samples 1 to 4) and AF2/BR2 (samples 5 to 8) were used. Templates: lanes 1 and 5, cDNA; lanes 2 and 6, no-RT control without reverse transcriptase; lanes 3 and 7, control without template; lanes 4 and 8, genomic DNA. M, molecular weight marker.

Generation and selection of G. oxydans mutants with a defect in PQQ biosynthesis. The pqqA gene of G. oxydans was disrupted by insertion of a kanamycin resistance gene, using the gene replacement vector pEX18Ap (24). As shown by colony PCR, the majority of the obtained kanamycin-resistant G. oxydans clones represented single-crossover mutants. To isolate double-crossover mutants, clones were screened on the basis of clearing zones on glucose-calcium carbonate agar (12). A PQQ-deficient pqqA::Kmr mutant, TH1, was selected from about 100 clones based on the absence of a clearing zone, indicating an inactive quinoprotein glucose dehydrogenase. Upon the addition of PQQ to the agar, the zone-clearing phenotype of TH1 could be restored to the appearance of G. oxydans wild-type colonies. The disruption of pqqA by the kanamycin cassette and the absence of the vector within the genome of mutant TH1 were confirmed by PCR.

To detect possible additional genes involved in PQQ biosynthesis in G. oxydans, Tn5 transposon mutagenesis was carried out. A PQQ-deficient transposon mutant, TH2, was selected from about 100 kanamycin-resistant clones based on the weak intensity of the clearing zone on glucose-calcium carbonate agar. As found for the pqqA::Kmr mutant, the zone-clearing phenotype of the transposon mutant TH2 could be restored to the appearance of wild-type colonies by the addition of PQQ.

The transposon insertion in mutant TH2 was located in the tldD gene (see below). To confirm that the phenotype of mutant TH2 was due to inactivation of tldD alone, a new mutant was generated by site-directed disruption of the tldD gene. For this purpose, the gene replacement vector pKmob18GII, which contains the gusA marker (29), was applied. Since pKmob18GII carries a kanamycin resistance gene, a gentamicin cassette was used for gene disruption. Double-crossover mutants were identified by blue-white screening as described previously (29). A colorless colony representing a double-crossover mutant (tldD::Gmr) was isolated and named TH3. The disruption of tldD by the gentamicin cassette and the absence of the vector within the genome of mutant TH3 were confirmed by PCR. Like in mutant TH2, only a faint clearing zone on glucose-calcium carbonate agar was visible with mutant TH3; a distinct zone of clearing could be restored by the addition of PQQ.

Sequence analysis of the tldD gene. The transposon Tn5 insertion site in the G. oxydans mutant TH2 was located by inverse PCR in a 1,464-bp open reading frame encoding a putative 51-kDa protein annotated as TldD (accession number AAW60873). The TldD protein family is listed as containing "predicted Zn-dependent proteases and their inactivated homologs" in the NCBI COG database. A BLASTP search and pairwise alignments using ClustalW revealed that the G. oxydans TldD protein showed 67% identity to TldD from Rhodospirillum rubrum ATCC 11170 (accession number YP_425658) as the best hit and 51% sequence identity to TldD from E. coli (accession number BAA07913). The E. coli TldD protein has been characterized by Allali and coworkers (2). tldD mutants accumulate a precursor of the peptide antibiotic MccB17, indicating that TldD is involved in processing or export of MccB17. TldD has also been found to participate in the degradation of CcdA, a component of the ccd poison-antidote system of the F plasmid. A more general physiological role in biodegradation of unstructured polypeptides has also been proposed for E. coli TldD (2).

Sequence analysis of the genetic environment of the G. oxydans tldD gene suggests that tldD is organized in a tricistronic operon (Fig. 1B). tldD is located downstream of genes encoding a hypothetical protein and a transaminase that are oriented in the same direction. The two intergenic regions within the putative operon are short and show a low promoter probability (promoter prediction score below 0.2), whereas the region upstream of the hypothetical protein gene most probably represents a promoter (promoter prediction score, 0.91). The putative tldD-containing operon is located 124 kb downstream of the pqqABCDE operon.

Properties of the pqqA knockout mutant. The pqqA::Kmr mutant TH1 was transferred several times on mannitol medium agar to dilute out residual PQQ derived from wild-type cells present during triparental mating. As shown, after two or three transfers on agar plates, mutant TH1 was unable to grow on complex medium agar containing D-mannitol, glycerol, or D-glucose as the sole energy source (Table 3). Growth was restored in all cases when PQQ (12 µM) was added to the agar. However, on complex medium agar containing D-gluconate, mutant TH1 grew without the need of PQQ addition.


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  		TABLE 3. PQQ production and growth of wild-type, mutant, and recombinant G. oxydans strains

Mutant TH1 could be cultured in minimal medium containing D-gluconate (2%) as the sole energy source; however, growth was very slow, and an apparent agglomeration of cells interfered with the determination of the optical density. Growth of TH1 was slightly improved when both D-gluconate and D-mannitol were added to the minimal medium, but a long lag phase was still observed. Mutant TH1 grew well in medium containing 2% D-gluconate, 2% D-mannitol, and low concentrations of tryptone (0.03%) and yeast extract (0.05%). This medium was therefore used for PQQ determination for TH1 cultures. No PQQ could be detected in TH1 supernatants, whereas supernatants of the wild type grown in the same complex medium contained about 50 ng PQQ/ml (Table 3), similar to wild-type supernatants of cultures grown in minimal medium. TH1 could be complemented by a plasmid carrying the pqqA gene, including the putative pqqA promoter region (pTB9031).

Membrane fractions of mutant TH1 were assayed for dye-linked dehydrogenase activities with D-mannitol, glycerol, and D-gluconate at pH 5 or D-glucose at pH 6 (Table 4). No significant activity was found for mutant membranes with D-mannitol, glycerol, or D-glucose. When mutant membranes where incubated with PQQ and cations (Mg2+ or Ca2+) to reconstitute quinoprotein apoenzymes to holoenzymes, mannitol and glycerol dehydrogenase activities were restored to levels comparable to those for wild-type membranes. In contrast, incubation of mutant membranes with PQQ resulted in only a minor restoration of glucose dehydrogenase activity: 9% (with Mg2+) or 17% (with Ca2+) of the activity of wild-type membranes was found. Prolonged incubation with PQQ and cations (30 min instead of the standard procedure with 10 min) did not enhance activity.


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  		TABLE 4. Dye-linked dehydrogenase activities in membrane fractions of G. oxydans wild type and mutants

In mutant membranes, dehydrogenase activity with D-mannitol, glycerol, or D-glucose was also absent when Tris-HCl at pH 8.75 was used as the buffer. Furthermore, screening with additional substrates showed that dehydrogenase activities with D-sorbitol, D-arabitol, meso-erythritol, D-fucose, D-mannose, and ethanol were found for wild-type membranes but not for pqqA mutant membranes. In contrast, with D-gluconate as the substrate and without prior addition of PQQ, mutant membranes exhibited a similar dehydrogenase activity to that of wild-type membranes. Incubation of mutant membranes with PQQ and cations did not increase the level of gluconate dehydrogenase activity.

Properties of the tldD knockout mutant. Unlike the pqqA knockout mutant TH1, the tldD knockout mutant TH2 was able to grow on complex medium agar containing D-mannitol, glycerol, or D-glucose as the sole energy source, albeit slowly (Table 3). Mutant TH2 exhibited similar tendencies to those of the pqqA knockout mutant. Thus, colony growth with D-mannitol, glycerol, or D-glucose was rather slow and could be restored to the appearance of the wild type by adding PQQ to the agar. Like the pqqA mutant, mutant TH2 grew on agar with D-gluconate as the sole energy source. Under these conditions, no difference in growth was detected whether PQQ was added or not. In supernatants of the tldD mutant TH2 grown in minimal medium to optical densities of 0.4 to 0.6, no PQQ was detected (Table 3). Mutant TH2 could be complemented either with plasmid pTB9046, carrying the tldD gene under control of the G. oxydans tufB promoter, or with plasmid pTB9045, carrying the putative tldD-containing operon, including its promoter region (tldD and two upstream genes, encoding a transaminase and a hypothetical protein) (Fig. 1B). TH2-pTB9045 and TH2-pTB9046 both produced PQQ in amounts comparable to that in the wild-type strain (Table 3). Membranes of mutant TH2 showed much lower dehydrogenase activities with D-mannitol, glycerol, and D-glucose than those of wild-type membranes (Table 4). As with the pqqA mutant TH1, mannitol and glycerol dehydrogenase activities in TH2 could be restored to levels similar to those of the wild type by incubation with PQQ and cations, whereas active glucose dehydrogenase could only be restored to 34% (with Mg2+) or 50% (with Ca2+). With D-gluconate as the substrate, TH2 membranes exhibited similar dehydrogenase activity to that of wild-type membranes.

The tldD::Gmr mutant TH3 that was generated subsequently by site-directed gene disruption (see above) exhibited similar growth behavior to that of the transposon mutant TH2. Like mutant TH2, mutant TH3 did not produce detectable amounts of PQQ, showed low dehydrogenase activity with D-mannitol, glycerol, and D-glucose, and could be complemented by the plasmid-encoded tldD gene under control of the G. oxydans tufB promoter or by the putative tldD-containing operon including its own promoter (data not shown).

PQQ overproduction in G. oxydans. PQQ was measured in supernatants of cultures grown to optical densities of 0.4 to 0.6. While supernatants of the G. oxydans wild-type strain contained about 45 ng PQQ per ml, a two- to threefold higher PQQ concentration was found for the recombinant strain harboring the plasmid-carried pqqA gene under control of its own promoter (pTB9031). In a recombinant strain harboring a plasmid bearing the complete pqqABCDE operon (pTB9028), PQQ production amounted to about 1,500 ng/ml and was therefore enhanced 30-fold compared to that in the wild type.

No further increase in PQQ synthesis was obtained in strains containing plasmids in which the G. oxydans pqqABCDE operon was cloned behind the E. coli tufB promoter or behind the G. oxydans tufB promoter. Instead, PQQ synthesis using these constructs was enhanced only about fivefold compared to that of the wild type. To test whether the tldD gene further enhances PQQ overproduction in G. oxydans, a plasmid carrying the tldD gene under control of the G. oxydans tufB promoter in addition to the pqqABCDE operon (pTB9055) was used. However, the recombinant strain containing this plasmid produced only 20 times more PQQ than the wild type (Table 3).


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DISCUSSION
 
The pqqABCDE(F) cluster present in different PQQ-synthesizing bacteria is generally referred to as an operon controlled by a single promoter upstream of pqqA. However, from sequence analysis it was suggested that in G. oxydans, pqqA carries its own promoter while the pqqBCDE genes are controlled by an additional weak promoter within the large pqqA-pqqB intergenic region (12). In contrast, our data demonstrate cotranscription of the pqqAB genes and the lack of detectable promoter activity of the pqqA-pqqB intergenic region, suggesting that the pqqA promoter represents the only promoter within the pqqABCDE cluster of G. oxydans 621H, a similar situation to that found for other PQQ-synthesizing bacteria. However, it cannot be excluded that the pqqA-pqqB intergenic region exhibits promoter activity under conditions not yet known.

So far, no gene replacement vector has been described for G. oxydans that allows easy selection of double-crossover mutants independent of the phenotype caused by the gene disruption. Therefore, in cases of phenotypes that are tedious to screen, the identification of double-crossover mutants can be difficult. In this study, the gene replacement vector pEX18Ap, which carries the Bacillus subtilis sacB gene as a counterselectable marker (24), was initially used for gene disruption. However, single-crossover mutants of G. oxydans having the vector integrated into the genome did grow on 5% or 10% sucrose, and the sacB gene strategy (15) could not be applied to isolate double-crossover mutants. Therefore, double-crossover mutants had to be identified based on their phenotype, i.e., the absence of quinoprotein glucose dehydrogenase activity (12). In the following experiments, the vector pKmobGII (29) was shown to be a useful tool for gene knockout in G. oxydans. Using blue-white screening based on the presence or absence of the vector-carried gusA gene, encoding a ß-glucuronidase, double-crossover mutants could be selected easily without the need for a screenable phenotype of the disrupted gene. As with Rhizobium meliloti 1021 and Agrobacterium tumefaciens A348 (29), blue colonies were observed 2 days after cell plating. pKmobGII was also used successfully to disrupt yet another G. oxydans gene encoding a PQQ-dependent dehydrogenase (data not shown).

The inability of the pqqA mutant to grow with D-mannitol, glycerol, or D-glucose as the sole energy source indicates that in G. oxydans 621H, PQQ is essential for growth with these substrates. In G. oxydans, D-mannitol, glycerol, and D-glucose are oxidized by membrane-bound quinoproteins that channel electrons into the respiratory chain. In the mutant lacking PQQ, these enzymes are inactive. Membrane fractions of the pqqA mutant did not exhibit any dye-dependent dehydrogenase activity with D-mannitol, glycerol, or D-glucose under the conditions used, indicating that in the cytoplasmic membrane, these substrates are oxidized exclusively by quinoproteins. Therefore, our findings demonstrate the vital role of quinoproteins in G. oxydans metabolism. In the genome of G. oxydans 621H, a total of 75 putative dehydrogenases of unknown function have been identified, 23 of which are predicted to be membrane bound (38). Three of these membrane-bound dehydrogenases share high homology with quinoproteins, while others belong to the flavin-containing enzyme families (38). Membrane-bound flavin-dependent enzymes carry out a similar function in respiratory energy generation to that of quinoproteins. The dye-linked assay used in this study measures both PQQ- and flavin-dependent enzymes. Therefore, our results suggest that G. oxydans 621H does not contain membrane-bound D-mannitol-, glycerol-, or D-glucose-oxidizing flavin-dependent dehydrogenases. Furthermore, the lack of dehydrogenase activity in pqqA mutant membranes with D-sorbitol, D-arabitol, meso-erythritol, D-fucose, D-mannose, and ethanol might suggest that in the membrane, these substrates are likewise oxidized exclusively by quinoproteins. D-Sorbitol, D-arabitol, and meso-erythritol are substrates of the quinoprotein glycerol dehydrogenase (34), while ethanol is oxidized by the quinohemoprotein alcohol dehydrogenase (33). D-Fucose and D-mannose are most probably substrates of the quinoprotein glucose dehydrogenase (19).

Whereas several quinoprotein apoenzymes, such as the glucose dehydrogenase of E. coli (25), can be converted readily into active holoenzymes in vitro, only a partial in vitro reconstitution by the addition of PQQ was reported for other quinoproteins (8). In membranes of the pqqA mutant, mannitol and glycerol dehydrogenase activities could be restored to levels comparable to those in wild-type membranes by the addition of PQQ. In contrast, glucose dehydrogenase activity could be reconstituted only to a minor degree in pqqA mutant membranes. This could be due to instability or poor reconstitutability of the apoenzyme with PQQ.

The pqqA mutant was still able to grow when D-gluconate was present in the medium. Consistently, mutant membranes showed gluconate dehydrogenase activity without prior addition of PQQ. In the G. oxydans 621H membrane, D-gluconate can be oxidized either by the quinoprotein glycerol dehydrogenase or by the flavin-dependent gluconate-2-dehydrogenase (11, 38), yielding 5-D-ketogluconate or 2-D-ketogluconate, respectively. Thus, in the mutant, respiratory energy generation could be driven by the flavin-dependent enzyme.

Despite the lack of quinoprotein glycerol dehydrogenase activity and thus the missing PQQ-dependent gluconate oxidation, the level of specific gluconate dehydrogenase activity in pqqA mutant membranes was at least the same as that in wild-type membranes. Possibly, under the conditions used, PQQ-dependent gluconate dehydrogenase activity is much lower than flavin-dependent (PQQ-independent) gluconate dehydrogenase activity, such that for measurements of total gluconate oxidation, a loss of PQQ-dependent activity is not apparent. Indeed, the quinoprotein glycerol dehydrogenases of G. oxydans strains IFO 3255 and IFO 3257 have much lower specific oxidizing activities with D-gluconate than with D-mannitol or glycerol (1, 49). Consistently, in our study, no decrease in gluconate dehydrogenase activity was detected in wild-type membranes after treatment with 10 mM EDTA, which converts the quinoprotein glycerol dehydrogenase into the apoform (1), whereas mannitol and glycerol dehydrogenase activities were completely abolished by the same treatment (data not shown). Furthermore, the slightly enhanced gluconate dehydrogenase activity in the pqqA mutant compared to the wild type might be due to some kind of activation or upregulation of the flavin-dependent gluconate-2-dehydrogenase, as was also found for a pqqE knockout mutant of G. oxydans ATCC 9937 (12), in which gluconate oxidizing activity was twofold higher than that in the wild-type strain (22).

Disruption of the tldD gene in G. oxydans led to a drop of PQQ excretion below the detection limit. Therefore, our study demonstrates that in addition to the pqqABCDE cluster, the tldD gene is involved in PQQ biosynthesis in G. oxydans 621H. The TldD protein of G. oxydans is related to E. coli TldD, a peptidase involved in processing of small peptides (2). In other PQQ-producing bacteria, the peptidase-like protein PqqF is required for PQQ production. An equivalent of the pqqF gene had not been recognized in G. oxydans so far. Therefore, we hypothesize that the G. oxydans TldD protein carries out a similar function to that of PqqF proteins in other PQQ-synthesizing species, i.e., processing the peptide precursor of PQQ. Sequence analysis showed that TldD of G. oxydans does not share significant similarity with known PqqF proteins; the maximum sequence identity was found for PqqF from Methylobacterium extorquens (12.7%). However, the known PqqF proteins all show very low sequence similarities; PqqF from Klebsiella pneumoniae and PqqF from Methylobacterium extorquens share only 10.3% sequence identity and belong to different subfamilies of peptidases. Thus, as already suggested (46), the different PQQ-synthesizing bacteria seem to use divergent proteins for processing of the peptide precursor. Furthermore, G. oxydans TldD is likely to also play a role in other, yet unknown processes, similar to E. coli TldD.

The ability of the tldD mutant to grow, albeit slowly, on D-mannitol, D-glucose, or glycerol as the sole energy source and the low but measurable dehydrogenase activities in membranes found with these substrates indicate that small amounts of PQQ, below the detection limit of the assay used, are produced even in the absence of TldD. Obviously, these small amounts of PQQ support a level of PQQ-dependent dehydrogenase activity that is sufficient for slow growth on D-mannitol, D-glucose, or glycerol. Consistent with our observation is the report that an E. coli strain transformed with the pqqABCDE genes of Klebsiella pneumoniae but missing the pqqF gene also produced small amounts of PQQ, although much less than the strain that contained all genes of the pqqABCDEF operon (52). It was suggested that other proteases present in E. coli could substitute, to a limited extent, for PqqF so that small amounts of PQQ were made even in the absence of the pqqF gene. By analogy, a protease present in G. oxydans could assume the function of TldD in the tldD mutant.

Transformation of the G. oxydans wild type with a plasmid containing the G. oxydans pqqABCDE cluster enhanced PQQ synthesis about 30-fold. However, at 1,500 ng PQQ per ml, the strain still produced less PQQ than wild-type strains of some methylotrophs, such as Methylobacterium organophilum XX, in which extracellular PQQ production of up to 2,600 ng/ml was found under optimized culture conditions (51). Due to the apparently weak activity of the pqqA promoter, the pqqABCDE genes were set under the control of the tufB promoter of G. oxydans or E. coli, the latter of which has already been used successfully in G. oxydans for the overproduction of 2-keto-L-gulonic acid (41). However, neither the use of the tufB promoter nor additional cloning of the tldD gene resulted in further enhancement of PQQ production. Nonetheless, our results demonstrate that the rather elaborate biosynthesis of PQQ, which requires the synthesis of a small peptide as a precursor of the cofactor, can be enhanced significantly by the simple overexpression of the pqqABCDE operon. Although involved in PQQ synthesis, the TldD protein does not seem to be limiting in this pathway.


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ACKNOWLEDGMENTS
 
We thank Irmgard Maue-Mohn and Viola Khodaverdi for excellent technical assistance and helpful discussions, Cagla Köken for technical assistance, Mustafa Elfari for participation in the initial design of the project, and Lorenz Adrian for helpful discussions. We also thank Ute Herrmann, Forschungszentrum Jülich, for information on the tufB promoter; Armin Ehrenreich and Marc Hoffmeister, Georg-August-University Göttingen, for information on the G. oxydans genome; Florian Jarosch and Christian Maasch, Noxxon Pharma AG, Berlin, for the RT protocol and helpful suggestions on RT and RT-PCR; Anke Becker, Universität Bielefeld, for the gift of plasmid pKmobGII; and Oliver Lenz and Bärbel Friedrich, Humboldt University, Berlin, for the gift of plasmid pCM130.

This project was carried out within the framework of the Competence Network Göttingen Genome Research on Bacteria (GenoMik) financed by the German Federal Ministry of Education and Research (BMBF).


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FOOTNOTES
 
* Corresponding author. Mailing address: FG Technische Biochemie, Sekr. GG1, TU Berlin, Seestr. 13, D-13353 Berlin, Germany. Phone: 49 30 31427509. Fax: 49 30 31427581. E-mail: Tina.Hoelscher{at}TU-berlin.de. Back

{triangledown} Published ahead of print on 25 August 2006. Back


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Journal of Bacteriology, November 2006, p. 7668-7676, Vol. 188, No. 21
0021-9193/06/$08.00+0     doi:10.1128/JB.01009-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.



This article has been cited by other articles:

  • Holscher, T., Weinert-Sepalage, D., Gorisch, H. (2007). Identification of membrane-bound quinoprotein inositol dehydrogenase in Gluconobacter oxydans ATCC 621H. Microbiology 153: 499-506 [Abstract] [Full Text]  

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