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Institute of Microbiology and Biotechnology, University of Ulm, D-89069 Ulm, Germany
Correspondence
Bernhard J. Eikmanns
bernhard.eikmanns{at}uni-ulm.de
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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-1,4 glucosyl linkages and a smaller number of -1,6 branching linkages. In most bacteria, glycogen accumulates in the stationary growth phase and under conditions of limited growth in the presence of an excess of carbon and energy (Preiss & Romeo, 1994
). Therefore, glycogen is generally assumed to be a storage compound serving as a carbon and energy reserve (reviewed by Preiss, 1984, 1996). Consistent with this assumption, some bacterial mutants deficient in glycogen synthesis show decreased survival under starvation conditions, when compared to the wild-type (Preiss, 1984). However, glycogen obviously is not essential for bacterial growth since many of these mutants grow as well as their parental strains (Preiss, 1984). In some bacteria, such as Bacillus subtilis and Streptomyces coelicolor, glycogen may play a role in sporulation and differentiation, respectively (Kiel et al., 1994; Martin et al., 1997), and in Mycobacterium smegmatis, glycogen has been proposed to serve as a carbon capacitor for glycolysis during exponential growth, in addition to its conventional storage role (Belanger & Hatfull, 1999). Furthermore, glycogen biosynthesis recently has been connected with osmotically regulated trehalose synthesis in Corynebacterium glutamicum (Tzvetkov et al., 2003; Wolf et al., 2003; Padilla et al., 2004b). This organism forms trehalose as a compatible solute under certain (hyper)osmotic stress conditions and additionally as an essential cell wall compound (Wolf et al., 2003; Puech et al., 2000; Argüelles, 2000; Daffé, 2005). In contrast to other bacteria, the osmotically induced trehalose synthesis in C. glutamicum has been shown to be mainly mediated by the TreYZ pathway (Wolf et al., 2003). This pathway uses (poly)maltodextrins, such as glycogen, as a substrate and thus, glycogen formation in C. glutamicum might be involved in osmoprotection of this organism.
The pathway of glycogen biosynthesis in bacteria proceeds by the action of three enzymes (Fig. 1), i.e. ADP-glucose (ADP-Glc) pyrophosphorylase (ATP : -D-glucose-1-phosphate adenyltransferase; EC 2.7.7.27), glycogen synthase (EC 2.4.1.21) and branching enzyme (EC 2.4.1.18), encoded by the glgC, glgA and glgB genes, respectively (Preiss, 1984, 1996). The key regulatory step of this pathway is the reaction of ADP-Glc pyrophosphorylase, which forms ADP-Glc and pyrophosphate from ATP and -D-glucose 1-phosphate. Most ADP-Glc pyrophosphorylases are allosterically controlled (positively and negatively) by various intermediates of the major carbon and energy metabolic pathways (Ballicora et al., 2003; Preiss, 1996, 1984). The genetic aspects of glycogen biosynthesis have been studied most comprehensively in Escherichia coli (reviewed by Preiss, 1996). In this organism the genes encoding the glycogen biosynthetic enzymes are organized in a cluster which includes the glgC, glgA and glgB genes and two genes involved in glycogen degradation, glgX and glgY (Romeo et al., 1998). The glgY gene is now designated glgP (Preiss, 1996) and encodes a glycogen phosphorylase (EC 2.4.1.1) which catalyses glycogen breakdown by removing glucose units from the nonreducing ends (Alonso-Casajús et al., 2006), whereas the glgX gene recently has been shown to encode an isoamylase-type debranching enzyme (EC 3.2.1–) with high specificity for hydrolysis of polysaccharide outer chains that have been recessed previously by glycogen phosphorylase (Dauvillée et al., 2005). The E. coli glg gene cluster is composed of two tandemly arranged operons (glgCAP and glgBX) and the transcription of these operons is subject to complex regulation including catabolite repression, stringent control and/or control by RpoS, KatF, GlgQ, GlgR and/or the carbon storage regulator CsrA (Baker et al., 2002; Romeo & Preiss, 1989; Yang et al., 1996; Preiss, 1996). Clustering of genes involved in glycogen metabolism has been also described for other bacteria, e.g. Bacillus stearothermophilus and B. subtilis (Takata et al., 1997; Kiel et al., 1994), Rhodobacter sphaeroides and R. capsulatus (Igarashi & Meyer, 2000), Mesorhizobium loti (Lepek et al., 2002) and Agrobacterium tumefaciens (Ugalde et al., 1998), although the precise genetic organization and transcriptional regulation obviously differs among these organisms.
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). The genome of this organism has recently been determined and annotated (GenBank accession nos NC_003450 and BX927147; Kalinowski et al., 2003; Ikeda & Nakagawa, 2003) and open reading frames putatively encoding several enzymes involved in glycogen metabolism were detected (Kalinowski et al., 2003). These genes include glgC (cg1269), glgA (cg1268), glgB (cg1381), glgE (cg1382), two glgP genes (cg1479 and cg2289) and a glgX gene (cg2310), the latter four possibly encoding a glucanase, two glycogen phosphorylases and a debranching enzyme, respectively. However, so far none of the respective enzyme activities have been been proven to be present in C. glutamicum and, with the exception of glgA, involvement of the genes and their products in glycogen synthesis or degradation has not been shown. In the course of studies on trehalose biosynthesis, Tzvetkov et al. (2003) recently found that inactivation of the chromosomal glgA gene led to the failure of sucrose-grown C. glutamicum cells to accumulate glycogen. Thus, although there is evidence for the presence of glycogen in C. glutamicum, knowledge of glycogen formation and degradation, the enzymes involved, their regulation, and the function of glycogen in C. glutamicum is meagre. In this communication we report on the glycogen content of C. glutamicum during growth in different media, its ADP-Glc pyrophosphorylase activity, the analysis, expression and inactivation of the glgC gene, and the significance of glycogen synthesis for amino acid production and for osmoprotection of C. glutamicum. |
METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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(Hanahan, 1983), C. glutamicum wild-type (WT; ATCC 13032 from the American Type Culture Collection) and its lysine-producing derivative C. glutamicum DM1730 (pycP458S homV59A lysCT311I zwfA243T) (Georgi et al., 2005; obtained from B. Bathe, Degussa AG, Halle-Künsebeck Germany). Plasmid pK19mobsacB (Schäfer et al., 1994) was employed for construction of the glgC mutants C. glutamicum WT-IMC and DM1730-IMC; plasmid pET2 (V
sicová et al., 1998) was used for transcriptional fusion experiments.
E. coli and all pre-cultures of C. glutamicum were grown aerobically in TY or LB complex medium (Sambrook & Russell, 2001) at 37 °C and 30 °C, respectively. For the main cultures of C. glutamicum, the cells of an overnight pre-culture were washed with 0.9 % (w/v) NaCl and inoculated into TY medium with or without 1 % (w/v) glucose or into minimal medium (Eikmanns et al., 1991) containing glucose, fructose, sucrose, acetate or lactate at concentrations indicated in the Results and Discussion section. C. glutamicum was grown aerobically at 30 °C as 50 ml cultures in 500 ml baffled Erlenmeyer flasks on a rotary shaker at 120 r.p.m. or as 250 ml cultures in a parallel fermentation system (see below). In standard glutamate fermentations, ethambutol (25 mg l–1) was added to the culture 2 h after inoculation, as described by Radmacher et al. (2005). The growth of E. coli and of C. glutamicum was monitored by measuring the OD600.
Parallel fermentations with C. glutamicum were performed at 30 °C in 450 ml glass fermenters (‘Fedbatch-pro system’ from DASGIP). The pH was maintained at 7.0 using a standard pH electrode (Broadley James Corporation) and addition of 2 M KOH and/or 2 M HCl. Foam development was controlled by manual injection of small amounts (about 100 µl) of silicon antifoam (Roth). The dissolved oxygen was measured online using a polarimetric oxygen electrode (Broadley James Corporation) and it was adjusted to 30 % of saturation during constant aeration with 2 vvm [30 l h–1] in a cascade by stirring at 150 to 1200 r.p.m.
Competition experiments between C. glutamicum WT and C. glutamicum WT-IMC were performed as follows. The OD600 of both pre-cultures was determined; the proportions of both pre-cultures in the common inoculum to give an OD600 of 1 in the main culture were calculated according to the ratios of C. glutamicum WT and C. glutamicum WT-IMC stated in the Results and Discussion section. The ratio of C. glutamicum WT and C. glutamicum WT-IMC in the culture broth during the competition experiments was monitored by live-cell counting on LB plates with and without kanamycin (25 µg ml–1).
DNA preparation, transformation and DNA manipulations.
Standard procedures were employed for plasmid isolation, for molecular cloning and transformation of E. coli DH5, and for electrophoresis (Sambrook & Russell, 2001). C. glutamicum chromosomal DNA was isolated according to Eikmanns et al. (1994). Plasmid DNA from E. coli was isolated using the method of Birnboim (1983). C. glutamicum WT and DM1730 were transformed by electroporation using the method of Tauch et al. (2002). The resulting transformants were selected on LB-BHIS agar plates containing kanamycin (25 µg ml–1). All restriction enzymes, T4 DNA ligase, calf intestine phosphatase and Taq polymerase were obtained from MBI-Fermentas or from Roche Diagnostics and used according to the instructions of the manufacturer.
RNA techniques.
Total RNA from C. glutamicum was isolated and purified as described by Schreiner et al. (2005). For Northern (RNA) hybridization, a digoxigenin-dUTP-labelled glgC-specific 0.5 kb DNA probe was generated by PCR with primers IM-glgC-for (5'-ACGCGTCGACACCACGTGTATCGCATGG-3') and IM-glgC-rev (5'-CGGGATCCTGTGGATTGGCCACTCAG-3'; BamHI restriction site underlined). For hybridization, total RNA from C. glutamicum WT was separated on an agarose gel containing 17 % (v/v) formaldehyde and transferred onto a nylon membrane (Eikmanns et al., 1994). Hybridization [at 50 °C, in the presence of 50 % (v/v) formamide], washing and detection were carried out using the Nucleic Acid Detection kit (Roche Diagnostics). The size marker was the 0.24–9.5 kb RNA ladder from GibcoBRL.
Cloning of the glgC promoter.
The glgC promoter fragment was amplified from chromosomal DNA of C. glutamicum WT by PCR with the primers PRAC-2-for (5'-ACGCGTCGACTGCACCCATGCAGTGAAC-3'; SalI site underlined) and PR-glgAC-rev (5'-CGGGATCCGAATCCGGAGTTCACCAG-3'; BamHI site underlined). The 437 bp PCR product, covering the region from 263 bp upstream to 174 bp downstream of the translational start codon, was digested with SalI and BamHI, ligated into SalI/BamHI-restricted plasmid pET2 and transformed into E. coli. The recombinant plasmid pET-PC was then isolated from E. coli and introduced into C. glutamicum by electroporation. The nucleotide sequence of the promoter fragment in plasmid pET-PC was verified by sequence analysis (MWG Biotech).
Construction of the glgC mutants of C. glutamicum.
The chromosomal glgC gene in C. glutamicum was disrupted by the method described by Schäfer et al. (1994). For this purpose, a 0.5 kb internal fragment of glgC was amplified by PCR with primers IM-glgC-for and IM-glgC-rev (see above), restricted with SalI and BamHI, and the resulting 0.37 kb DNA fragment was inserted into plasmid pK19mobsacB, which is nonreplicative in C. glutamicum. The resulting plasmid pK19IMC was transformed into C. glutamicum WT and DM1730; transformants were obtained by selection on medium containing kanamycin (15 µg ml–1). Kanamycin resistance indicated integration of pK19IMC into the chromosomal glgC gene via recombination. To confirm integration, we performed a Southern blot analysis, essentially as described by Reinscheid et al. (1999). BamHI-restricted chromosomal DNA from C. glutamicum WT, and from putative glgC integration mutants C. glutamicum WT-IMC and DM1730-IMC, was hybridized to a digoxigenin-dUTP-labelled 0.37 kb internal glgC probe (prepared from plasmid pK19IMC), resulting in two signals, at 9.2 kb and 7.1 kb, with the DNAs from C. glutamicum WT-IMC and DM1730-IMC and one signal, at 10.2 kb, with DNA from C. glutamicum WT. These sizes were expected for the glgC integration mutants and the WT strain, respectively.
Analysis of intracellular glycogen.
The glycogen content of C. glutamicum cells was determined either enzymically (Parrou & Francois, 1997) or by TLC essentially as described by Seibold et al. (2006). For the enzymic assays, 5 ml samples of the culture were harvested and the cells were washed twice with TN buffer (50 mM Tris, 50 mM NaCl; pH 6.3) and once with resuspension buffer (40 mM potassium acetate; pH 4.2). The cells then were suspended in resuspension buffer (total volume 1 ml) and transferred to 2 ml screw-top Eppendorf tubes filled with 250 mg acid-washed glass beads (150–212 µm; Sigma-Aldrich). After inactivation of cell-bound glycosidic activity by incubation at 95 °C for 5 min, the cells were disrupted using a Ribolyser (Hybaid) three times without cooling at maximum speed of 6.5 for 45 s. The cell debris and glass beads were separated from the supernatant by centrifugation (13 000 g, 20 min). Each sample was divided into two 100 µl aliquots (assays A and B), and 2 µl amyloglucosidase (10 mg ml–1; Roche Diagnostics) was added to assay A; assay B was used as a reference. Both assays were incubated with shaking for 2 h at 57 °C. The glucose concentration was then measured in both control (assay B; without amyloglucosidase) and test (assay A; with amyloglucosidase) tubes as described below. The amount of glucose determined in the control assay B was subtracted from the amount of glucose determined in assay A. The cell dry weight of C. glutamicum was calculated according to the OD600; an OD600 of 1 corresponded to 0.25 g l–1 for C. glutamicum (Börmann et al., 1992).
TLC was used to distinguish between glycogen and other (lower molecular mass) carbohydrates that might serve as substrates for amyloglucosidase. Aliquots (10 ml) of the culture were withdrawn, and the cells were pelleted by centrifugation for 5 min at 13 000 g and washed twice with TN buffer following a washing step with water. The cell pellet was resuspended to a total volume of 1 ml with water and transferred into a 2 ml screw-top Eppendorf tube filled with 250 mg acid washed glass beads. Enzyme activity was inactivated, cells were disrupted and cell debris and glass beads were removed by centrifugation as described above. Sample preparation, application, separation and visualization of the carbohydrates on the TLC plate (by spraying with sulphuric acid solution and successive charring) were performed as previously described (Seibold et al., 2006). Carbohydrates were identified by comparison to the migration of authentic standards (glucose, maltose, maltotriose, maltoheptaose and glycogen; all except glucose obtained from Sigma Aldrich).
Quantification of glucose and of amino acids.
For analysis of the glucose or amino acid concentrations in the culture broth, 1 ml of the culture was withdrawn and centrifuged (13 000 g, 10 min, 4 °C) and the supernatant was used for the determination. The amino acid concentrations were determined as described by Schrumpf et al. (1991). Glucose was determined enzymically using hexokinase/glucose-6-phosphate dehydrogenase (Roche Diagnostics) and spectrophotometric quantification (at 340 nm) of the NADPH formed.
Enzyme assays.
ADP-Glc pyrophosphorylase activity was determined by measuring the formation of glucose 1-phosphate from ADP-Glc and pyrophosphate similar to the method described by Takata et al. (1997). C. glutamicum cells were harvested, washed twice in 0.1 M Tris/HCl, pH 8, 20 mM KCl, 5 mM MgSO4 and resuspended in 0.75 ml of the same buffer containing 0.1 mM EDTA, 2 mM DDT and 10 % (v/v) glycerol. The cell suspension was transferred to 2 ml screw-cap vials together with 250 mg glass beads and subjected five times for 30 s to mechanical disruption with a Ribolyser at 4 °C, with intermittent cooling on ice for 5 min. After disruption, the glass beads and cellular debris were removed by centrifugation (10 000 g, 4 °C, 15 min). Afterwards the supernatant was centrifuged (45 000 g, 1.5 h) to remove the membrane fraction. Carbohydrates such as maltodextrins, serving as a substrate for background glucose 1-phosphate formation, were removed by use of a HiTrapDesalting column (Amersham Bioscience) equilibrated with 0.1 M Tris/HCl, pH 8, 25 mM MgCl2 on an Äkta Purifier System (Amersham Bioscience). Up to 200 µl of the enzyme preparation was added to a total volume of 500 µl of reaction mixture containing 0.1 M Tris/HCl, pH 8, 25 mM MgCl2, 2 mM sodium pyrophosphate, 1.5 mM iodonitrotetrazolium chloride (INT), 5 U phosphoglucomutase (from rabbit muscle; Roche Diagnostics), 5 U glucose-6-phosphate dehydrogenase (from Leuconostoc sp.; Roche Diagnostics) and 5 U diaphorase (from pig heart, Roche Diagnostics). The reaction was initiated at 30 °C by addition of 5 mM ADP-Glc and the change of absorption at 492 nm due to the reduction of INT was monitored in a spectrophotometer (Ultrospec 2100 pro, Amersham Bioscience) for 3 min. As a control, the change in absorption at 492 nm in the absence of ADP-glucose was measured and subtracted from the experimental values. One unit is defined as the amount of enzyme that produces 1 µmol glucose 1-phosphate in 1 min.
To determine chloramphenicol acetyltransferase (CAT) activity, crude extracts were prepared as described above, except that the cells were washed twice in 0.1 M Tris/HCl, pH 7.8, and resuspended in the same buffer containing 10 mM MgCl2 and 1 mM EDTA. The specific CAT activity was determined as described by Schreiner et al. (2005). The biuret method (Gornall et al., 1949) with bovine serum albumin as the standard was used to determine protein concentrations.
Computational analysis.
MFold (Zuker, 2003) software was used for the calculation of the 
G0' value (free energy under standard conditions) of the glgC terminator structure. The protein and DNA sequences were analysed using the ExPASy proteomics server (Gasteiger et al., 2003) and CLUSTALW (Thompson et al., 1994). The SWISSPROT accession number for C. glutamicum ADP-Glc pyrophosphorylase is Q8NRD4. Additional ADP-Glc pyrophosphorylase sequences used for comparison with this polypeptide and their respective accession numbers are as follows: Mycobacterium tuberculosis (P64241), S. coelicolor (P72394), E. coli (POA6V1) and A. tumefaciens (Q8V8L5).
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RESULTS AND DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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). With 5 % (w/v) glucose, the cells grew to an OD600 of about 60. Cells of all four cultures showed rapid glycogen accumulation in the early exponential growth phase and degradation of the polymer before they entered the stationary phase, i.e. when the carbon source became limiting. However, the maximal intracellular glycogen concentration varied in the cells grown on the different substrates (Fig. 2b). Whereas C. glutamicum in minimal medium with glucose or sucrose accumulated up to 90 mg glycogen per g dry weight, the cells cultured in minimal medium with fructose as single carbon source accumulated only about 40 mg glycogen per g dry weight. The lower glycogen accumulation on this substrate may be due to different entry points of the substrates into the central metabolism, i.e. glucose 6-phosphate (the direct precursor of glucose 1-phosphate) in the case of glucose and sucrose and fructose 1,6-phosphate (which has to be dephosphorylated and isomerized to glucose 6-phosphate) in the case of fructose (Yokota & Lindley, 2005). It should be noted that the maximal glycogen content of the C. glutamicum cells was about 35 % lower when they were grown in media containing only 1 % (w/v) glucose instead of 2 %. Although increasing the initial glucose concentration to 5 % (w/v) led to slightly higher glycogen concentrations in the early stationary phase (about 45 mg per g dry weight at after 9 h), it did not lead to higher intracellular glycogen concentrations or to a significantly different glycogen profile in the course of growth.
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ADP-Glc pyrophosphorylase activity in C. glutamicum
The specific activity of ADP-Glc pyrophosphorylase was determined in cell extracts of C. glutamicum WT grown in TY complex medium and in minimal medium containing glucose, potassium acetate or sodium lactate [all at 1 % (w/v)] as the carbon source and harvested at the early exponential growth phase (3 h after inoculation). The highest specific activity [6.0 mU (mg protein)–1] was found in cells grown in minimal medium containing glucose. The activity was about 20-fold lower when the cells were grown in TY complex medium or in minimal medium containing acetate or lactate [0.2–0.4 mU (mg protein)–1]. The dependency of ADP-Glc pyrophosphorylase activity on the growth phase was elucidated by measuring the specific activity in extracts from C. glutamicum WT in minimal glucose medium harvested at the early, mid and late exponential phase and at the stationary phase. The highest specific activity was found when the cells were harvested in the early exponential phase [6.0 mU (mg protein)–1]; slightly lower activities were found in cells at the mid and late exponential phase [5.2 mU (mg protein)–1] and at the stationary phase [3.8 mU (mg protein)–1]. These results suggest that ADP-Glc pyrophosphorylase activity in C. glutamicum is severely regulated by the carbon source in the growth medium and only very weakly regulated by the growth phase. However, it should be stated that the specific ADP-Glc pyrophosphorylase activity did not correlate with the cellular glycogen content observed in the course of growth. This result indicates that glycogen synthesis in C. glutamicum is controlled either by another biosynthetic enzyme (e.g. glycogen synthase or branching enzyme) or by activity regulation of ADP-Glc pyrophosphorylase by (a) so far unidentified effector(s). Here it is noteworthy to mention that all known bacterial ADP-Glc pyrophosphorylases, with the exception of those in B. stearothermophilus and B. subtilis, are allosterically controlled by small effector molecules, such as metabolites of the glycolytic pathways (as activators) and AMP, ADP and inorganic phosphate (as inhibitors) (reviewed by Ballicora et al., 2003). The probability of allosteric regulation of the C. glutamicum ADP-Glc pyrophosphorylase is strengthened by the identification of amino acid residues known to be important for the regulation of the corresponding E. coli enzyme (see below).
Analysis of the glgC gene and its chromosomal organization
The C. glutamicum cg1269 gene (identical to NCgl1073) was sequenced in the course of the determination of the genome sequence (accession nos NC_003450 and BX927147) and has been designated ADP-Glc pyrophosphorylase gene glgC (Kalinowski et al., 2003; Ikeda & Nakagawa, 2003). It has a length of 1218 bp and the predicted gene product (accession no. Q8NRD4) consists of 405 amino acids with a molecular mass of 43 861 Da. The glgC gene is preceded by a probable ribosome-binding site (AAGGG) and followed by a region of dyad symmetry similar to rho-independent transcription terminators [centred 18 bp downstream of the TAA stop codon; G0'=–14.7 kcal mol–1 (–61.5 kJ mol–1) at 25 °C]. This result indicates transcriptional termination downstream of the glgC gene. The glgC gene is located upstream of and in the opposite orientation to cg1268, which putatively encodes a glycogen synthase and is therefore annotated as glgA. Tzvetkov et al. (2003) inactivated this gene in C. glutamicum and found the mutant to be unable to accumulate glycogen. Downstream of, and also in the opposite orientation to glgC, there is a gene (cg1270) annotated as ‘probable O-methyltransferase’ (Kalinowski et al., 2003). Except for a putative fructose hydrolase gene (cg1267) directly downstream of glgA, no other genes involved in glycogen or carbohydrate metabolism were identified near glgC and glgA. Putative glgB (cg1381), glgP (cg1479 and cg2289) and glgX (cg2310) genes (encoding glycogen synthetic and degradation enzymes) have been identified elsewhere in the C. glutamicum genome (Kalinowski et al., 2003) and thus, unlike the clustering of five glg genes (glgC, glgA, glgB, glgP and glgX) in other bacteria (Igarashi & Meyer, 2000; Preiss, 1996), the C. glutamicum glg genes (or their homologues) obviously are scattered within the genome.
Database analysis and alignment studies with the C. glutamicum protein encoded by glgC revealed significant similarity of the C. glutamicum protein to the functionally well-characterized ADP-Glc pyrophosphorylase from E. coli (Preiss & Romeo, 1994; 40 % identity) and other Gram-negatives (such as A. tumefaciens; 42 %), from S. coelicolor (62 %), and to putative ADP-Glc pyrophosphorylases from other high-G+C Gram-positive bacteria, such as Corynebcterium efficiens (91 %), C. diphtheriae (79 %), C. jeikeum (76 %) and M. tuberculosis (70 %). An alignment of the deduced amino acid sequence of the C. glutamicum glgC gene product with the respective sequences of the E. coli enzyme and of some other representative bacteria (Gram-positives and Gram-negatives) is shown in Fig. 3. All the enzymes aligned contain highly conserved stretches, including the E. coli ADP-Glc pyrophosphorylase regions known to carry amino acids involved in catalysis or regulation (reviewed by Ballicora et al., 2003). Asp142 and Arg32 in the E. coli ADP-Glc pyrophosphorylase, proposed to be involved in catalysis, correspond to Asp127 and Arg19, respectively, in the C. glutamicum enzyme. Lys195 and Tyr114, known to have specific roles in the binding of substrates (glucose 1-phosphate and ATP, respectively) in the E. coli enzyme, correspond to the C. glutamicum Lys180 and Tyr105 residues, and Pro295 and Gly336, important for the regulation of the E. coli enzyme to Pro284 and Gly323, respectively, in the C. glutamicum polypeptide. A sequence motif ‘Arg-Ala-Lys-Pro-Ala-Val’, which is present in all ADP-Glc pyrophosphorylases (residues 27–32 in the C. glutamicum enzyme), has also been shown to be important for activator binding (Greene et al., 1996).
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; Preiss, 1996).
To confirm the presence of a glgC-specific promoter and to investigate transcriptional regulation of the glgC gene, a transcriptional fusion between the putative glgC promoter region and the promoterless chloramphenicol acetyltransferase (CAT) gene was constructed in the promoter probe vector pET2. The resulting plasmid, pET-PC, was transformed into C. glutamicum WT and CAT activity was determined in the plasmid-carrying strain during early exponential growth in TY medium and in minimal medium containing glucose, acetate or lactate [each at 1 % (w/v)]. Whereas C. glutamicum carrying the host plasmid pET2 showed no CAT activity [<0.01 U (mg protein)–1] on either medium, C. glutamicum (pET-PC) showed activity of 0.10–0.16 U (mg protein)–1 when grown in TY medium with or without glucose or in minimal medium containing acetate or lactate. In cells grown in minimal medium containing glucose, CAT activity was about threefold higher [0.40–0.45 U (mg protein)–1]. In the course of growth (from 3 to 24 h after inoculation), none of these activities changed significantly. These results confirm the presence of a promoter immediately upstream of the glgC gene and indicate weak transcriptional control of the glgC gene by the carbon source in the growth medium. In E. coli, the expression of the glgC gene is subject to positive regulation by cAMP, the cAMP receptor protein CRP and by ppGpp (reviewed by Preiss, 1996). However, in C. glutamicum there is no indication of a carbon catabolite repression mechanism involving cAMP/CRP or involving a catabolite control protein (CcpA) and a catabolite-responsive element (CRE), the typical regulatory elements in carbon catabolite repression of low-G+C Gram-positive bacteria (Hueck et al., 1994; Stülke & Hillen, 2000). Also, there is no indication for an involvement of the stringent response since a C. glutamicum rel gene deletion mutant, unable to synthesize (p)ppGpp, showed unaltered glgC expression when compared to the wild-type strain (Brockmann-Gretza & Kalinowski, 2006).
The threefold higher promoter activity in the cells grown in glucose medium instead of acetate or lactate medium did not correspond to the 20-fold higher ADP-Glc pyrophosphorylase activities observed in glucose-grown cells when compared to cells grown on the other carbon sources (see above). This result suggests that the observed carbon-source-dependent regulation of ADP-Glc pyrophosphorylase activity is only to a small extent due to transcriptional regulation of the glgC gene and mainly due to post-transcriptional control at the RNA or at the enzyme level. In E. coli glgC gene expression has been shown to be post-transcriptionally regulated by the CsrA protein, by acceleration of glg transcript degradation (Liu et al., 1995). Taken together, the data suggest that glgC in C. glutamicum is under transcriptional and post-transcriptional control; however, the nature and the mechanisms of glgC expression control need to be investigated in future studies.
Inactivation of the chromosomal glgC gene and characterization of the mutant
To study whether C. glutamicum requires the glgC gene for growth, for ADP-Glc pyrophosphorylase activity and for intracellular glycogen accumulation, the chromosomal glgC gene was inactivated by integration mutagenesis (see Methods). The resulting strain, C. glutamicum WT-IMC, was then analysed in comparison to the WT C. glutamicum. Both C. glutamicum WT and C. glutamicum WT-IMC grew equally well (measured by doubling time and final optical density) in LB and TY complex medium and in minimal medium containing glucose, fructose, sucrose, acetate or lactate (not shown). Thus, an intact glgC gene is not required for growth in any of these media. In contrast, inactivation of glgC led to a loss of the vast majority of ADP-Glc pyrophosphorylase activity [0.4 mU (mg protein)–1 in the mutant instead of 6 mU (mg protein)–1 in the WT strain] and to a complete loss of intracellular glycogen in all media tested (data not shown). These results indicate (i) that glgC in fact encodes an ADP-Glc phosphorylase and (ii) that a functional glgC gene is required for intracellular glycogen accumulation in C. glutamicum. The low residual activity in the ADP-Glc pyrophosphorylase assay with C. glutamicum WT-IMC cells might be due to an ADP-Glc pyrophosphatase (EC 3.6.1.21) activity, which releases sugar 1-phosphate from ADP-sugar. The cg1607 gene, encoding a protein with similarity (25 % identity) to the E. coli ADP-Glc pyrophosphatase (Moreno-Bruna et al., 2001) and carrying the typical ‘nudix’ motif (Mildvan et al., 2005), has been annotated as a pyrophosphohydrolase in the C. glutamicum genome (Kalinowski et al., 2003).
We also studied growth, substrate consumption and glycogen content of C. glutamicum WT and the glgC mutant WT-IMC in glucose-minimal medium in a controlled parallel fermentation system providing a constant pH of 7.0 and a constant oxygen concentration of 30 % saturation. C. glutamicum WT and WT-IMC grew at nearly identical growth rates of 0.36 h–1 and reached approximately the same final OD600 of 35 (Fig. 4a). At the beginning of the exponential growth phase, glucose was consumed considerably faster by C. glutamicum WT than by the glgC mutant (Fig. 4a). This faster glucose consumption of the WT strain was paralleled by extensive glycogen formation (Fig. 4b). The glycogen content of the WT was maximal in the early exponential growth phase and then sharply decreased. Twelve hours after inoculation, the glucose in both cultures was exhausted (Fig. 4a), and quantitative enzymic analysis of the glycogen content as well as qualitative analysis of intracellular carbohydrates via TLC (Fig. 4c) showed almost complete degradation of the intracellular glycogen in C. glutamicum WT. In contrast, the mutant strain C. glutamicum WT-IMC formed no glycogen at any time tested (Fig. 4b, c).
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Direct competition experiments in minimal glucose medium with C. glutamicum WT and the WT-IMC mutant were performed to test for a possible growth advantage of the WT strain. Both strains were inoculated together at different ratios (100 %, 80 %, 50 % and 20 % C. glutamicum WT-IMC), consecutively grown in minimal medium with 2 % (w/v) glucose, and the percentage of the mutant was determined in the course of the cultures. No changes were observed in the ratio between the WT and the mutant strain WT-IMC in the course of three consecutive cultures. This result shows that the WT strain is not able to outcompete the glgC mutant in minimal glucose medium and thus suggests that glycogen formation is not advantageous under the conditions tested.
In summary, the results of the characterization of the mutant indicate (i) that the glgC gene in C. glutamicum in fact encodes a protein with ADP-Glc pyrophosphorylase activity, (ii) that ADP-Glc pyrophosphorylase of C. glutamicum is involved in intracellular glycogen formation, (iii) that intracellular glycogen formation is not essential for growth of C. glutamicum in any medium tested and (iv) that the inability to form intracellular glycogen is no disadvantage for C. glutamicum under all conditions tested here.
Significance of glycogen synthesis for amino acid production
As C. glutamicum is used for the industrial production of L-glutamate and L-lysine from glucose, it was interesting to investigate the influence of the inactivation of glycogen synthesis on amino acid formation. In standard glutamate fermentations in minimal medium containing 2 % glucose and using ethambutol as the trigger C. glutamicum WT and WT-IMC accumulated about the same concentration of glutamate (22.1 and 19.0 mM, respectively). To analyse the significance of glycogen formation for L-lysine production, we constructed a glgC-negative derivative of the lysine producer C. glutamicum DM1730. The resulting mutant C. glutamicum DM1730-IMC accumulated approximately the same concentration of L-lysine as the parental strain DM1730 (36.7 and 38.9 mM, respectively). These results indicate that the absence of glycogen synthesis does not significantly affect L-glutamate and L-lysine production by C. glutamicum.
Role of glycogen synthesis in osmoprotection of C. glutamicum
For C. glutamicum, adaptation to hyperosmotic stress caused by changes in the osmolality of the environment is necessary for survival in its natural habitat, soil. The organism counteracts shifts to high osmolality by the synthesis of compatible solutes, such as glutamate, proline and trehalose (Wolf et al., 2003). The TreYZ pathway, one of two major pathways leading to trehalose formation in C. glutamicum (Wolf et al., 2003; Tzvetkov et al., 2003), starts from linear maltodextrins, which are intermediates in glycogen synthesis. To investigate the influence of glycogen synthesis on growth during hyperosmotic stress, C. glutamicum WT and mutant WT-IMC were cultivated in minimal medium with glucose and 750 mM NaCl. Compared to the WT strain, the mutant showed a lag phase of about 2 h and then commenced to grow with a rate of about 0.22 h–1. Both strains grew to approximately the same final OD600, 30.5 and 29.8, respectively (Fig. 5a). In the early stages of growth, glucose consumption by the WT strain was slightly higher than by the mutant; however, glucose was used up by both strains within 14 h (Fig. 5a). No glycogen was formed throughout growth by the mutant strain WT-IMC (Fig. 5b). The glycogen formation profile of the WT, however, showed differences when compared to that during growth of this strain in minimal medium without NaCl (Fig. 4b and 5b
, respectively). The maximal glycogen level of about 50 mg glucose (g dry weight)–1 was observed in the middle of the exponential growth phase and the degradation of glycogen was significantly slower (Fig. 5b).
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). Thus, the availability of maltodextrins (due to the synthesis of glycogen) and the constitutive presence of TreY and TreZ guarantee an immediate response to osmotic stress, whereas in the absence of either glycogen synthesis or the TreYZ pathway, the enzymes for the alternative trehalose synthetic pathways (OtsA and OtsB) have to be formed, and thus the trehalose response of C. glutamicum to hyperosmotic conditions takes some time. That the OtsAB pathway principally can substitute the TreYZ pathway for trehalose synthesis became clear from studies with single and/or double mutants defective in either or both of the pathways (Wolf et al., 2003; Tzvetkov et al., 2003) and from treYZ and otsAB overexpression studies, which aimed at improvement of microbiological trehalose production with C. glutamicum (Padilla et al., 2004a; Carpinelli et al., 2006).
Taken together, the results indicate that glycogen accumulation in C. glutamicum is favourable for fast adaptation to hyperosmotic stress under the conditions tested. Since hyperosmotic stress is also a factor in large-scale amino acid fermentations, glycogen accumulation might also be of advantage for the industrial amino acid production process. It should be noted here, however, that the significance of trehalose as a compatible solute in C. glutamicum depends on environmental conditions (Guillouet & Engasser, 1995; Wolf et al., 2003; Tzvetkov et al., 2003) and also that other compatible solutes (such as proline) are involved in the response of C. glutamicum to hyperosmotic conditions. Thus, the relevance of glycogen synthesis for fast osmoprotection might be restricted to conditions in which trehalose represents the main compatible solute.
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ACKNOWLEDGEMENTS |
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Edited by: M. Hecker
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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