Role of central metabolism in the osmoadaptation of the halophilic bacterium chromohalobacter salexigens

Role of Central Metabolism in the Osmoadaptation of the Halophilic Bacterium Chromohalobacter salexigens * □ S Received for publication, March 18, 2013, and in revised form, April 8, 2013 Published, JBC Papers in Press, April 24, 2013, DOI 10.1074/jbc.M113.470567 José M. Pastor ‡1,2 , Vicente Bernal ‡1,3 , Manuel Salvador §2 , Montserrat Argandoña §4 , Carmen Vargas § , Laszlo Csonka ¶5 , Ángel Sevilla ‡6 , José L. Iborra ‡ , Joaquín J. Nieto § , and Manuel Cánovas ‡7 From the ‡ Departamento de Bioquímica y Biología Molecular B e Inmunología. Facultad de Química, Campus Regional de Excelencia Internacional “Campus Mare Nostrum,” Universidad de Murcia, 30100 Murcia, Spain, the § Departamento de Microbiología y Parasitología, Universidad de Sevilla, 41012 Seville, Spain, and the ¶ Department of Biological Sciences, Purdue University, West Lafayette, Indiana 47907-2064 Background: Chromohalobacter salexigens synthesizes and accumulates ectoines. Results: High ratio of the anaplerotic and catabolic fluxes involved in ectoines synthesis supports high biosynthetic fluxes at high salinity and leads to metabolite overflow at low salinity. Conclusion: Evolution optimized the metabolism of C. salexigens to support high production of ectoines. Significance: Metabolic adaptations in a compatible solute-accumulating halophile are described for the first time. Bacterial osmoadaptation involves the cytoplasmic accumulation of compatible solutes to counteract extracellular osmolarity. The halophilic and highly halotolerant bacterium Chromohalobacter salexigens is able to grow up to 3 MNaCl in a minimal medium due to the de novo synthesis of ectoines. This is an osmoregulated pathway that burdens central metabolic routes by quantitatively drawing off TCA cycle intermediaries. Consequently, metabolism in C. salexigens has adapted to support this biosynthetic route. Metabolism of C. salexigens is more efficient at high salinity than at low salinity, as reflected by lower glucose consumption, lower metabolite overflow, and higher biomass yield. At low salinity, by-products (mainly gluconate, pyruvate, and acetate) accumulate extracellularly. Using [113 C]-, [213 C]-, [613 C]-, and [U13 C 6 ]glucose as carbon sources, we were able to determine the main central metabolic pathways involved in ectoines biosynthesis from glucose. C. salexigens uses the Entner-Doudoroff pathway rather than the standard glycolytic pathway for glucose catabolism, and anaplerotic activity is high to replenish the TCA cycle with the intermediaries withdrawn for ectoines biosynthesis. Metabolic flux ratios at low and high salinity were similar, revealing a certain metabolic rigidity, probably due to its specialization to support high biosynthetic fluxes and partially explaining why metabolic yields are so highly affected by salinity. This work represents an important contribution to the elucidation of specific metabolic adaptations in compatible solute-accumulating halophilic bacteria. Halophilic microorganisms demand relatively high salt concentrations to grow. Because of the diversity of environments where they can thrive, their physiology is widely varied. The metabolic diversity of halophilic and halotolerant microorganisms is conditioned by the adaptation to specific environmental niches (especially evident in the case of alkalophilic, methanotrophic, or thermophilic halophilic bacteria) and also by osmoadaptation mechanisms that these microorganisms developed to cope with salinity (1–3). As a consequence, the preferred metabolic pathways used to assimilate carbon sources are also diverse. Strategies of osmoadaptation can be roughly classified in two main types. The “salt-in” strategy, which consists of the accumulation of K ⫹ and Cl ⫺ in the cytoplasm of the cells, is used by extremely halophilic aerobic archaea, halophilic fermentative bacteria, and the extremely halophilic bacterium Salinibacter ruber (4–6). The “organic solutes-in” strategy, which involves the accumulation of organic “compatible” solutes, is used by a larger variety of organisms, including all mesophilic bacteria, halophilic algae, halophilic methanogenic archaea, and halotolerant and halophilic aerobic bacteria (7). Among the halophilic eubacteria that use the organic solutes-in strategy, strict aerobiosis is more frequent because compatible solute synthesis is energetically and metabolically a very demanding process (4). Compatible solutes belong to a few chemical families: sugars (sucrose and trehalose), polyols (glycerol, glucosylglycerol, mannosylglycerol, and arabitol, among others), amino acids (glutamine and derivatives, proline, alanine), quaternary amines (betaines and choline), and ectoines (ectoine and ␤ -hydroxyectoine). Ectoine is one of the most widely distributed *This work was supported in part by Fondo Europeo de Desarrollo Regional funds, Ministerio de Ciencia e Innovación (Spain) Projects BIO2008-0450201, BIO2011-29233-C02-01, and BIO2011-22833, Junta de Andalucía (Spain) Grant P08-CVI-03724, and Spanish National Network on Extremophilic Microorganisms Grant BIO2011-12879-E. □ S This article contains supplemental Materials and Methods, Tables S1–S4, Figs. S1–S8, and additional references. 1 Both authors contributed equally to this work. 2 Recipient of Formación de Profesorado Universitario and Formación de Personal Investigador fellowships from MICINN (Spain). 3 Recipient of a post-doctoral contract from Universidad de Murcia (Programa Propio). To whom correspondence may be addressed: Grupo de Bioenergía, Dirección de Tecnología, Centro de Tecnología de Repsol, Carretera A-5, Km 18, 28935 Móstoles-Madrid, Spain. Tel.: 34-868-887393; Fax: 34-868-884148; E-mail: [email protected]. 4 Recipient of a post-doctoral contract from Junta de Andalucía. 5 Supported by National Science Foundation Award IOS-1054977. 6 Recipient of a post-doctoral contract from the Programa Juan de la Cierva (Spain). 7 To whom correspondence may be addressed. Tel.: 34-868-887393; Fax: 34-868-884148; E-mail: [email protected]. THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 288, NO. 24, pp. 17769–17781, June 14, 2013 © 2013 by The American Society for Biochemistry and Molecular Biology, Inc. Published in the U.S.A. JUNE 14, 2013•VOLUME 288•NUMBER 24 JOURNAL OF BIOLOGICAL CHEMISTRY 17769 at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from compatible solutes. Hydroxyectoine is often synthesized at lower amounts together with ectoine by many ectoine-producing species (8). When present in the medium, either these compounds or their precursors can be taken up from the environment. When cells are growing in media lacking compatible solutes, these compounds can only be accumulated by de novo synthesis (9). Chromohalobacter salexigens DSM 3043 (formerly Halomonas elongata DSM 3043) is a halophilic ␥ -proteobacterium of the family Halomonadaceae (10). It has one of the widest salinity ranges for growth found in nature (10, 11). In C. salexigens, osmoadaptation is mainly achieved by the accumulation of ectoine and hydroxyectoine (designated together as “ectoines”) (9). The biosynthetic pathway for ectoines was established in H. elongata and C. salexigens (11–15). The availability of the genomic sequence of C. salexigens (16) presents an opportunity for the understanding of the characteristic metabolic features of this halophile and their relevance for osmoadaptation. During active growth, metabolite fluxes are substantially directed toward biosynthetic pathways, draining intermediaries of central metabolism. Therefore, the perfect coupling and efficiency of the metabolic pathways linking carbon sources to the end products (in our case, compatible solutes) is crucial. There are two major pathways for the catabolism of sugars to pyruvate: glycolysis (Embden-Meyerhof pathway) (EM) 8 and the Entner-Doudoroff (ED) pathway (17). The physiological relevance of these pathways for the growth of C. salexigens is not known (16, 18, 19). The synthesis of ectoines consumes acetyl-CoA, which is produced by oxidative decarboxylation of pyruvate, and oxaloacetate (OAA), which is an intermediate in the TCA cycle and has to be replenished by anaplerotic pathways (16). The flux ratios between anaplerotic and catabolic pathways are highly relevant for metabolic adaptation. Metabolic studies in halophilic and halotolerant bacteria are scarce, especially if a focus on the biosynthesis of compatible solutes is sought. In this work, we gained new insights into the role of central metabolism in the osmoadaptation of C. salexigens. Using isotope label tracing, we analyzed the pathways for glucose catabolism and how central metabolism copes with the high metabolic burden caused by ectoines biosynthesis. Our results show how the adaptations developed by this bacterium affect metabolic efficiency at different salinities and represent a step further in the understanding of the physiology of halophilic and halotolerant bacteria. MATERIALS AND METHODS Bacterial Strains and Cultures C. salexigens CHR61, a rifampicin-resistant spontaneous mutant of C. salexigens DSM 3043 T , was used throughout this study. For ectoine production and for characterization of metabolic pathways, the strain was grown in minimal medium M63 (pH 7.2) containing 16.3 g/liter KH 2 PO 4 , 4.2 g/liter KOH, 2 g/liter (NH 4 ) 2 SO 4 , 39.5 mg/liter MgSO 4 䡠7H 2 O, 0.5 mg/liter FeSO 4 䡠7H 2 O. M63 was supplemented with 35.0, 43.8, 146.0, or 175.2 g/liter NaCl (corresponding to 0.6, 0.75, 2.5, or 3 M). As a carbon source, 20 mMglucose was used. Aerobic 100-ml batch cultures were grown in 0.5-liter flasks at 37 °C on a rotary shaker at 210 rpm. Cultures were started from frozen 20% glycerol stocks. Precultures were grown in SW-2 medium (containing 2% (w/v), or 0.3 M, total salts) composed of 15.6 g/liter NaCl, 4.07 g/liter MgSO 4 䡠7H 2 O, 2.6 g/liter MgCl 2 䡠6H 2 O, 0.4 g/liter KCl, 67 mg/liter CaCl 2 䡠2H 2 O, 47 mg/liter NaBr, and 13 mg/liter NaHCO 3 (20). M63 cultures were inoculated to an initial absorbance (A 600 ) of 0.025 with an exponential phase preculture grown overnight in SW-2 medium. Glycerol stocks, solid culture media, and precultures were supplemented with rifampicin to a final concentration of 25 ␮ g/ml. Analytical Procedures Cell Growth—To measure cell concentration, cells were resuspended in a NaCl solution (0.6 to 3.0 M), and absorbance was measured at 600 nm (Novaspec Plus Visible Spectrophotometer, Amersham Biosciences). A 600 and gram of dry cell weight (g DCW ) were correlated for the strain used, according to the following empirical equations: g DCW /liter ⫽0.597䡠A 600 (for 0.6 Mgrown cultures), g DCW /liter ⫽0.557䡠A 600 (for 0.75 M grown cultures), and g DCW /liter ⫽0.532䡠A 600 (for 2.5 and 3 M grown cultures). Determination of Extracellular Organic Acids—Extracellular organic acids were determined by ion exchange chromatography. Acetate was analyzed in a Shimadzu LC-10 HPLC instrument (Shimadzu Scientific Instruments, Columbia, MD), equipped with differential refractive index and diode array (UV) detectors (Shimadzu Scientific Instruments, Columbia, MD). A cation exchange HPX-87H column (Bio-Rad) was used for the separation of organic acids. The mobile phase was 5 mM H 2 SO 4 at a 0.5 ml䡠min ⫺1 flow rate and 45 °C. Gluconate (m/z 195), pyruvate (m/z87), lactate (m/z89), and citrate (m/z191) were measured using HPLC-MS. Analysis was performed with an Agilent 1200 series HPLC instrument (Agilent Technologies, Santa Clara, CA) coupled to an Agilent 6120 single quadrupole mass spectrometer with orthogonal electrospray ionization source. The mass spectrometer was operated in the negative electrospray ionization mode, using the SCAN mode at a range of m/z50–300, whereas the selected ion monitoring mode was used for the m/zof each compound. The ion spray voltage was set at 3500 V. Nitrogen with a flux of 12 liters/min was used as the sheath gas (40 p.s.i.) and the auxiliary gas. The ion transfer capillary was heated to 350 °C. The fragmentation voltage was set at 70 V. Separation was carried out on an injection volume of 10 ␮ l using the same column and conditions as the previous method, substituting 5 mMH 2 SO 4 with 0.1% formic acid. Data were acquired by a PC using the Agilent Chemstation software. Isotopic Labeling Studies and NMR Spectroscopy—For the labeling experiments, cells were grown in 100 ml of M63 medium in the presence of isotopically labeled glucose (Cortec8 The abbreviations used are: EM, Embden-Meyerhof; ED, Entner-Doudoroff; OAA, oxaloacetate; Gdh, glutamate dehydrogenase; Pc, pyruvate carboxylase; Ppc, phosphoenolpyruvate carboxylase; Icdh, isocitrate dehydrogenase; Cs, citrate synthase; Pfk, 6-phosphofructokinase; Gdh, glutamate dehydrogenase; PEP, phosphoenolpyruvate; Pdh, pyruvate dehydrogenase. Central Metabolism and Osmoadaptation in C. salexigens 17770 JOURNAL OF BIOLOGICAL CHEMISTRY VOLUME 288•NUMBER 24•JUNE 14, 2013 at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from Net, Voisins-Le Bretonneux, France). Isotopically labeled substrates used were 100% [113 C]-, [213 C]-, [613 C]glucose, or 20% [U13 C 6 ]glucose. Cultures were harvested in the mid to late exponential phase (A 600 1.5 to 3), and cells were separated from supernatants by centrifugation (16,000 ⫻g, 15 min, 4 °C). Supernatants were concentrated by lyophilization, redissolved in 1 ml of deuterated methanol (Sigma) and used for the identification of extracellular by-products. 13 C nuclear magnetic resonance ( 13 C NMR) spectra were recorded on a Brucker AV200 spectrometer at 200 MHz and 25 °C, with a relaxation time of 1.5 s. Signals for pyruvate, gluconate, and acetate were assigned by comparison with previously published chemical shifts (Spectral Database for Organic Compounds) and confirmed by comparison with 13 C NMR spectra of pure compounds. Compatible solutes (ectoines, glutamate, and trehalose) and membrane lipids were extracted from the cell pellets by a variation of the protocol described by García-Estepa et al. (12). The aqueous phase was used for the analysis of compatible solutes, and the chloroform phase for membrane lipids. Spectra were recorded at 25 °C using Brucker AV400 and Brucker AV600 spectrometers at 400 and 600 MHz, respectively, and a relaxation time of 3 s. Peak areas were integrated for relative quantification. Spectrophotometric Determination of Glucose and Ammonia Consumption—Glucose was assayed by a glucose (hexokinase) assay kit (GAHK20, Sigma). Ammonium was assayed by an enzymatic assay kit (11112732035, from R-Biopharm, Darmstadt, Germany). Kits were used according to the recommendations of the manufacturers. Measurements were performed in a 96-well microplate reader Synergy HT (Bio-Tek, Winooski, VT). Enzyme Assays Enzyme assays were optimized for the conditions, media, and the microorganisms used in this work. Measurements were carried out in a 96-well microplate reader Synergy HT (Bio-Tek, Winooski, VT). A unit of enzyme activity was defined as micromoles of substrate consumed or product formed per min and was normalized to milligrams of protein (units䡠mg ⫺1 ). In each case, reactor bulk samples were withdrawn, and cells were centrifuged (16,000 ⫻g, 15 min, 4 °C) and resuspended in 65 mMphosphate buffer (pH 7.5). Cells were sonicated on ice with a 3-mm diameter probe using a Vibra Cell VC 375 ultrasonic processor (Sonics Materials, Danbury, CT) and centrifuged (16,000 ⫻g, 20 min, 4 °C). The supernatant (cell-free extract) was used for subsequent activity measurements. Protein concentration in cell-free extracts was determined by the bicinchoninic acid (BCA) method (BCA Protein Assay kit, Pierce). The protocols for the assessment of the activity of 6-phosphofructokinase (Pfk) (21), glucose-6-phosphate dehydrogenase (21), citrate synthase (Cs) (21), NADP ⫹ /NAD ⫹ -dependent isocitrate dehydrogenase (Icdh) (22), pyruvate carboxylase (Pc) (23), phosphoenolpyruvate carboxylase (Ppc) (23), isocitrate lyase (22), malic enzyme (24), aspartate aminotransferase (25), NADPH and NADH-dependent glutamate dehydrogenase (Gdh) (26) were optimized for C. salexigens (see supplemental material). In Silico Analysis of Protein Homology The completely sequenced and annotated genome of C. salexigens is available on line. However, the annotations have been made by automated homology studies of ORFs from many microorganisms, and there may be some incorrect assignments or gaps in the information. To analyze in detail the ORF assignments of the metabolic pathways more relevant for this work, we compared the metabolic reconstruction made by Ates et al. (18) with information available at genome sequence-based databases, such as the Kyoto Encyclopedia of Genes and Genomes (KEGG) (27), and MetaCyc (28), which are supported by experimental data. Gene and protein sequences were compared using BLAST (29). Analyses of domains (Conserved Domain Database, www.ncbi.nlm.nih.gov), protein localization and topology (Signal P 4.0 server), and genomic context were also performed. Prediction of the Fates of Isotopic Labels [113 C]-, [213 C]-, and [613 C]glucose were selected specifically for interrogating the relative importance of different pathways of central metabolism, as described previously (30, 31). The patterns of incorporation of the isotopic label from glucose into pyruvate and acetyl-CoA via glycolysis or the Entner-Doudoroff pathway and then into ectoines were predicted. RESULTS High Salinity Favors Biomass and Ectoine Production by C. salexigens To study the effect of salinity on the metabolism of C. salexigens, the production of biomass and ectoines was determined in cultures grown with glucose as the sole carbon source at 0.6, 0.75, 2.5, and 3 MNaCl. Maximum biomass production increased with salinity up to 2.5 MNaCl (Table 1), whereas TABLE 1 Growth and production of ectoines of C. salexigens at different NaCl concentrations Cultures were grown at 37 °C in M63 minimal medium with 20 mMglucose and 30 mMammonium as the sole carbon and nitrogen sources (see “Materials and Methods” for details). NaCl concentration in medium Maximum biomass Y Ect/X a Ectoines production rate Hydroxyectoine/ectoine ratio g DCW 䡠liter ⫺1 mmol䡠g DCW ⫺1 mmol䡠(g DCW 䡠h) ⫺1 0.6 M1.40 ⫾0.07 0.18 ⫾0.01 0.049 ⫾0.003 0.048 ⫾0.097 0.75 M1.76 ⫾0.03 0.45 ⫾0.01 0.127 ⫾0.004 0.106 ⫾0.030 2.5 M2.48 ⫾0.02 1.10 ⫾0.05 0.174 ⫾0.008 0.507 ⫾0.037 3M2.03 ⫾0.26 1.68 ⫾0.09 0.159 ⫾0.009 0.423 ⫾0.036 a Stoichiometric coefficient of ectoines production. Y Ect/X and ectoines production rates were determined in the exponential phase of cultures. Maximum biomass and hydroxyectoine to ectoine ratio were obtained in the early stationary phase. Central Metabolism and Osmoadaptation in C. salexigens JUNE 14, 2013•VOLUME 288•NUMBER 24 JOURNAL OF BIOLOGICAL CHEMISTRY 17771 at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from growth rate was optimal in minimal medium M63 with 0.75 M NaCl (Fig. 1) (9, 12). We reported previously that the intracellular content of ectoines increases with salinity (12), in agreement with the role of ectoines in osmoprotection. The stoichiometric coefficients of ectoines synthesis from biomass (Y Ect/X ) showed that total ectoines content was directly proportional to salinity (Table 1 and Fig. 2A). However, ectoines production rate was maximal at 2.5 MNaCl (Table 1). The hydroxyectoine to ectoine ratio increased with salinity up to 2.5 MNaCl (Table 1 and Fig. 2A), and an inverse correlation was observed between the relative content of ectoines and proteins with increasing salt concentration (Fig. 2B). Consumption of Carbon and Nitrogen Sources As the biosynthesis of ectoines occurs at the expense of central metabolic intermediates, it is expected to significantly burden metabolism. To assess salinity-dependent metabolic changes, nutrients and by-products were monitored at different salinities. Glucose is the favorite carbon source for C. salexigens (32). The stoichiometric coefficient of glucose consumption was highly affected by salinity, being highest at 0.6 MNaCl and decreasing by 75% at 3 MNaCl (Fig. 2C). Ammonium consumption was quite different, because the stoichiometric coefficient of ammonium uptake remained practically unaltered regardless of salinity (Fig. 2C). This is a remarkable finding, because, in principle, the high production of ectoines at high salinity should lead to a higher demand of the nitrogen source. To better understand the overall alterations of metabolism at different salinities, we calculated the ammonium to glucose consumption molar ratio. This parameter increased with salinity up to 2.5 MNaCl (Fig. 2C), and a positive correlation between the specific ectoines production rate, and the ammonium to glucose consumption ratios was observed (Fig. 2, Aand B). Overall, our findings show that the slow down in growth and metabolism at high salt concentrations favor biomass production, which is in agreement with the previously observed effect of salt concentrations above 1.5 MNaCl (32). This underscores that the higher efficiencies of carbon and nitrogen metabolism at high salinity are the consequences of the specialization to cope with a highly demanding environment. Quantification of Organic Acids Excreted by C. salexigens, Overflow Metabolism The presence of gluconate, acetate, pyruvate, and minor amounts of lactate in supernatants of cultures grown at 0.6 and 0.75 MNaCl suggested a possible overflow metabolism. The consumption of glucose and ammonium and production and reutilization of organic acids were determined at three different salinities. At any salt concentration, glucose was the growthlimiting nutrient, because its depletion marked the entrance into stationary phase. In contrast, around 15 mMammonium was still present in the medium at the end of growth of each culture (data not shown). At 0.6 and 0.75 MNaCl, acetate was produced during the exponential phase of growth (Fig. 1, Aand B), although at high salinity (2.5 MNaCl) extracellular acetate was almost undetectable (Fig. 1C). The specific rate of production of pyruvate production during exponential growth decreased with salt concentrations (Table 2). Acetate and pyruvate were re-assimilated once glucose was totally depleted, in contrast with gluconate, which accumulated in the culture medium during early exponential phase of growth and was consumed along with glucose in the mid-to-late exponential phase (Fig. 1). As described in Table 2, gluconate, pyruvate, and acetate were the major excreted products. Production rates of the two latter conditions inversely correlated to the salt concentration and the biomass yield. These findings suggest an increased metabolic efficiency at high salinity. The presence of these compounds in cultures grown suggested a possible overflow metabolism at low salinity. This FIGURE 1. Formation of biomass (black circles) and extracellular concentrations of glucose (black squares), gluconate (white circles), pyruvate (white squares), and acetate (white triangles) of batch cultures grown in 20 mMglucose minimal medium with 0.6 MNaCl (A), 0.75 MNaCl (B), and 2.5 MNaCl (C). Central Metabolism and Osmoadaptation in C. salexigens 17772 JOURNAL OF BIOLOGICAL CHEMISTRY VOLUME 288•NUMBER 24•JUNE 14, 2013 at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from could arise from a limited catabolism of glucose, leading to a higher excretion of by-products due to the reduced demand for ectoines. In Silico and in Vitro Analyses of Metabolic Pathways Related to the Synthesis of Precursors of Ectoines In Silico Analysis Accumulation of compatible solutes at high salinity imposes a biosynthetic burden on cells. The above results reflect that osmoadaptation in C. salexigens has implications on metabolic performance. C. salexigens genome has been automatically annotated, and preliminary analyses (16, 19) and a first genome-based metabolic reconstruction (18) have been published. To further understand the interplay between osmoadaptation and metabolism, we critically assessed a number of routes in this metabolic network related to central metabolism, with emphasis on the pathways leading to precursors of ectoines, as well as to metabolites found in supernatants. For this purpose, we performed the following: (i) homology studies using the information of related microorganisms such as Pseudomonads, Enterobacteria, and halotolerant bacteria available in metabolic databases such as MetaCyc and KEGG (27, 28); (ii) analysis of conserved domains, protein localization, and genomic context; (iii) growth experiments with D-glucono-1,5-lactone, D-gluconate, and 2-keto-D-gluconate as carbon sources, and (iv) review of literature data. Genome analysis revealed interesting metabolic features. The ED pathway, which is a route for the catabolism of glucose to pyruvate (33), could be operative in C. salexigens (supplemental Fig. S1). Typically, microorganisms using the ED pathway lack glycolytic enzyme(s), such as Pfk (34, 35). In fact, unambiguous annotation of the gene encoding Pfk in C. salexigens has been difficult (19). There are five ORFs annotated in the JGI website (genome.ornl.gov) as potential phosphofructokinases, although they have low homology with bona fide Pfks (supplemental Fig. S2 and supplemental Table S1). There is also a putative pyrophosphate-dependent phosphofructokinase (Csal1534). The reaction catalyzed by this enzyme is readily reversible (36). However, the gene for this enzyme from Propionibacterium freudenreichii can only complement fructose1,6-bisphosphatase (fbp) mutations but not phosphofructokinase (pfkA/pfkB) mutations in Escherichia coli (37), indicating that it functions in the gluconeogenic direction but not in the glycolytic direction. Significantly, the analysis of the C. salexigens genome also failed to identify a clear-cut representative of a fructose bisphosphatase (19), raising the possibility that Csal1534, which has been annotated as Ppi-Pfk, may be a fructose bisphosphatase. Our in silico analysis suggests that oxidation of D-glucose to D-gluconate through D-glucono-1,5-lactone could occur both in the periplasm and the cytoplasm. This agrees with our finding of the early accumulation of D-gluconate in the growth medium. C. salexigens was able to grow with D-glucono-1,5lactone, D-gluconate, and 2-keto-D-gluconate as the sole carbon source (supplemental Fig. S4), and a variant of the 2-keto-gluconate loop described in Pseudomonas (38) was predicted to be functional in C. salexigens (Fig. 3 and supplemental Fig. S1). FIGURE 2. Effect of NaCl concentration on production of ectoines and consumption of carbon and nitrogen sources in C. salexigens.A, cellular contents of ectoine and hydroxyectoine. B, relative content of proteins and ectoines, expressed as percentage of total pool of ectoines plus proteins. The sum of these pools was approximately constant throughout all conditions tested (0.423 ⫾0.055 g/g CDW ). C, stoichiometric coefficient of glucose (dark bars) and ammonium consumption (light bars). Molar ratio of ammonium to glucose utilization is denoted by circles. Cultures were grown at 37 °C in M63 minimal medium with 20 mMglucose and 30 mMammonium as the sole carbon and nitrogen sources, respectively. See the text for details. TABLE 2 Specific consumption/production rates of the main extracellular metabolites Cultures were grown in glucose/M63 minimal medium. See under “Materials and Methods” for details. All rates were calculated in the early exponential phase of growth and (except for lactate production) are expressed in mmol䡠g CDW ⫺1 䡠h ⫺1 . [NaCl] Glucose Ammonium Gluconate Pyruvate Lactate a Acetate M 0.6 14.28 ⫾1.28 3.73 ⫾2.87 0.87 ⫾0.31 2.25 ⫾1.20 0.00 ⫾0.00 0.43 ⫾0.17 0.75 11.73 ⫾3.24 6.46 ⫾0.75 1.14 ⫾0.38 1.66 ⫾0.51 13.2 ⫾9.5 0.53 ⫾0.20 2.5 2.1 ⫾0.18 2.48 ⫾0.40 0.97 ⫾0.47 0.30 ⫾0.17 0.00 ⫾0.00 0.02 ⫾0.01 a Lactate production rates are in ␮ mol䡠g CDW ⫺1 䡠h ⫺1 . Central Metabolism and Osmoadaptation in C. salexigens JUNE 14, 2013•VOLUME 288•NUMBER 24 JOURNAL OF BIOLOGICAL CHEMISTRY 17773 at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from C. salexigens possesses genes specifying putative Pc, Ppc, and OAA decarboxylase (supplemental Table S1). These enzymes interconvert pyruvate, phosphoenolpyruvate, and OAA and could have a role in supporting high ectoine biosynthetic fluxes by replenishing OAA needed for the TCA cycle (Fig. 3). The observed production of acetate is difficult to explain in the light of the current genome annotation and knowledge of metabolic pathways. Acetate metabolism in C. salexigens is quite different from that of Pseudomonas and E. coli. The main route of acetate production in P. aeruginosa, E. coli, and related species is the phosphotransacetylase acetate kinase pathway (39), which is not present in C. salexigens. Acetate production in the latter bacterium could involve the formation of the high energy intermediate acetyl phosphate through an alternative route. Csal1010 is annotated as a soluble pyruvate oxidase. This FAD-dependent enzyme decarboxylates pyruvate, producing acetate (28). The catabolism of ectoines also yields acetate (40); in fact, continuous synthesis and degradation of ectoines at low salinity could explain the higher acetate overflow. Regarding nitrogen metabolism, there is one copy of genes for alanine aminotransferase, L-alanine dehydrogenase, glutamate synthase, and glutamate dehydrogenase in the C. salexigens genome. The enzymes specified by these genes are responsible for reductive transfer of ammonium to 2-ketoglutarate to generate glutamate (41, 42), which acts as the major ammonium donor in the cell. There are two putative aspartate aminotransferases, which catalyze the reversible transfer of the amino group from glutamate to oxaloacetate, rendering aspartate and 2-ketoglutarate. This is a key enzyme as it links the TCA cycle with the first enzyme of the ectoines synthesis pathway (aspartokinase). C. salexigens has only one aspartokinase catalyzing the formation of aspartyl phosphate, which is a common metabolic intermediate in the biosynthesis of ectoines and aspartate family of amino acids (8, 19). For a complete description of the in silico analysis, see supplemental Table S1 and Figs. S1–S4. In Vitro Analysis Activities of selected enzymes were determined in vitro. The enzymes assayed belonged to four main groups as follows: (i) glucose/gluconate metabolism; (ii) TCA cycle; (iii) anaplerotic and gluconeogenic reactions, and (iv) nitrogen metabolism. Enzyme activities were determined in the mid-exponential phase cultures at different salinities (Table 3). FIGURE 3. Scheme of the central metabolism and synthesis of ectoines in C. salexigens based on the annotated genome. Pathways leading from glucose to 6-P-gluconate (6PGln) are proposed on the comparison of the in silico analysis of C. salexigens and P. putida. Abbreviations used are as follows: 2-KGlcn, 2-ketogluconate; 2-Kglu, 2-ketoglutarate; 6PGlcn, 6-phospho-D-gluconate; 6PKGlcn, 6-phospho-2-keto-D-gluconate; AcCoA, acetyl-coenzyme A; AcP, acetyl phosphate; Ala,L-alanine; Amm, ammonium; Asp,L-aspartate; Asp-P,L-aspartyl phosphate; G3P, glyceraldehyde 3-phosphate; Glc,Dglucose; Glc6P,D-glucose 6-phosphate; Glcn,D-gluconate; Glcnlac,D-gluconolactone; Gln,L-glutamine; Gox, glyoxylate; Glu,L-glutamate; Ict,D-isocitrate; KDGlcn6P, 2-keto-3-deoxy-Dgluconate-6-phosphate; Lac,D-lactate; Lys,Llysine; Mal, L-malate; Met,L-methionine; NADA, N- ␥ -acetyl-L-2,4-diaminobutyrate; OAA, oxaloacetate; PEP, phosphoenolpyruvate; Pyr, pyruvate; Suc, succinate; Thr, L-threonine; Acs, acetyl-coenzyme A synthetase; ActP, acetate permease; AcyP, acetyl phosphate phosphatase; AlaAT, alanine aminotransferase; Ald, alanine dehydrogenase; AspAT, aspartate aminotransferase; AspK, aspartate kinase; Cs, citrate synthase; EctA, diaminobutyrate acetyltransferase; Gad, gluconate dehydrogenase; Gdhq, glucose dehydrogenase; Gdh, glutamate dehydrogenase; Glk, glucokinase; Gls, glutamate synthase; GnuK, gluconokinase; Icl, isocitrate lyase; KguD, 2-keto-6-phosphogluconate reductase; KguK, 2-ketogluconate kinase; Ldh, lactate dehydrogenase; Mae, malic enzyme; Mqo, malate-quinone oxidorreductase; Ms, malate synthase; Oad, oxaloacetate decarboxylase; Pc, pyruvate carboxylase; Pdh, pyruvate dehydrogenase; Pox, pyruvate oxidase; Ppc, phosphoenolpyruvate carboxylase; Pyk, pyruvate kinase; Zwf, glucose-6-phosphate dehydrogenase. Dashed arrows are used for conversions that require more than one enzymatic step. Central Metabolism and Osmoadaptation in C. salexigens 17774 JOURNAL OF BIOLOGICAL CHEMISTRY VOLUME 288•NUMBER 24•JUNE 14, 2013 at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from Glucose Metabolic Enzymes, Glucose-6-phosphate Dehydrogenase and Pfk—Glucose-6-phosphate dehydrogenase interconnects the EM, ED, and pentose phosphate pathways (43) and is considered as a major route for NADPH production for biosynthesis and redox homeostasis (Table 3). Significantly, we were unable to detect Pfk activity above background in cell-free extracts (Table 3 and supplemental Tables S1 and S2 and Figs. S1–S3). This fact along with the lack of unequivocal assignment for Pfk suggests that, like Pseudomonads and some other aerobic genera (44), C. salexigens uses the ED pathway for glucose catabolism, whereas the operation of functional glycolysis remains uncertain (Fig. 3). TCA Cycle Enzymes, Cs and Icdh—In addition to their important role for the supply of metabolic energy, TCA cycle intermediates are key building blocks for the synthesis of biomass and ectoines (Fig. 3). Two activities of the TCA cycle were determined, Cs and Icdh. Two Icdh-encoding genes are annotated (Csal0525 and Csal1434), which differ in cofactor specificity; however, only NADP ⫹ -dependent Icdh activity was detected. Regardless of salinity, Icdh activity was 8–15-fold higher than that of Cs (Table 3). Anabolic and Anaplerotic Pathways Enzymes, Ppc, Pc, Isocitrate Lyase, and Malic Enzyme—Anaplerotic pathways are essential to replenish the OAA in the TCA cycle that is withdrawn for the production of ectoines (Fig. 3). Both Ppc and Pc activities were detected, the latter being 8–10-fold higher than the former. Interestingly, the Pc and Ppc activities measured in cells grown at 2.5 MNaCl were 2-fold higher than observed at low salinities (Table 3). The activity of the anaplerotic enzymes isocitrate lyase (isocitrate lyase and glyoxylate shunt) and malic enzyme (malic enzyme, gluconeogenesis) (45) was low compared with other activities analyzed. This is in agreement with what has been described in glucose-grown E. coli and Pseudomonas aeruginosa (46, 47). Nitrogen Metabolism Enzymes, Aspartate Aminotransferase and Glutamate Dehydrogenase—Glutamate dehydrogenase (Gdh) along with glutamine synthetase and glutamate synthase are the routes of ammonium assimilation in bacteria. Both NADHand NADPH-dependent Gdh activities were detected in C. salexigens extracts. Only the NADH-Gdh enzyme was predicted from the genomic sequence of C. salexigens (Csal1340) suggesting that it may not discriminate between the two pyridine nucleotides. Transaminases, catalyzing the transfer of the amino group between amino acids, are involved in amino acid synthesis. Aspartate aminotransferase activity was high, which should not be surprising considering that this activity must account for the synthesis of ectoines and all amino acids from the aspartate family (Fig. 3). Enzyme Activities and in Vivo Fluxes—Enzyme activities determined in vitro can be viewed as estimates of flux through a given route. Intracellular carbon fluxes can be roughly estimated from glucose uptake rates. When compared with Cs and Pc activities, the following is evident: (i) Cs activity was similar at all three salt concentrations, whereas Pc activity was higher at 2.5 MNaCl (Table 3), and (ii) the glucose uptake rate was 5–30-fold higher than Cs and Pc activities (the difference being higher at low salinity, supplemental Table S3). Although this is a rough approximation, these facts are in accord with our data on the accumulation of extracellular metabolites (Table 2) explaining why cells divert a significant part of the metabolized glucose to pyruvate and acetate overflow and suggesting that the TCA cycle might be limited by the low Cs activity. In addition, the high Pc activity indicated that OAA was actively synthesized from pyruvate. Tracing Ectoines Labeling from Glucose Finally, to assess the distribution of fluxes of central metabolism, the biosynthetic pathways for ectoines production were traced by isotopic labeling with [113 C]-, [613 C]-, and [213 C]glucose at low and high salinity. All possible isotopomers derived from these precursors via the EM and ED pathways, anaplerosis and the TCA cycle, were predicted for pyruvate, PEP, OAA, acetyl-CoA, and ectoines, based on the annotated genome (supplemental Fig. S5 and Tables S3 and S4). Ectoines Labeling in [113 C]- and [613 C]Glucose-grown Cultures—The analysis of the labeling of ectoines with [113 C]- and [613 C]glucose provides a way to estimate the EM to ED flux ratio. In cells grown with [113 C]glucose, the COOH and C6 of ectoines were predominantly labeled, regardless of the salt concentration, and the labeling of the other carbons was close to the natural abundance of the 13 C isotope (Fig. 4). The labeling of COOH can be explained by use of the ED pathway. Surprisingly, C6 of ectoines contained substantially higher amount of 13 C than would be predicted from natural abundance. We can account for this excess labeling of C6 of ectoines by postulating that 13 CO 2 generated by decarboxylation of [113 C]pyruvate is reincorporated efficiently by the carboxylation of either Pc or Ppc. The enrichment of label at the COOH of ectoines allowed us to estimate that over 95% of glucose used for ectoines synthesis was metabolized through the ED pathway and Pc. Label scrambling due to other pathways such as the pentose phosphate pathways could explain minor label enrichment at other positions. Also, the existence of a functional but minor glycolysis cannot be firmly dismissed. TABLE 3 Enzyme activities in crude extracts of exponential phase of batch cultures grown in glucose/M63 containing 0.6, 0.75, and 2.5 MNaCl All values are given in milliunits䡠(mg protein) ⫺1 . Data are the averages of eight determinations (four independent cultures, assayed in duplicate). NM means not measured. Zwf is glucose-6-phosphate dehydrogenase; Icl, isocitrate lyase; Mae, malic enzyme; AspAT, aspartate aminotransferase. Enzyme activity NaCl concentration 0.6 M0.75 M2.5 M Gluconate metabolism Zwf 39.1 ⫾15.5 55.7 ⫾27.4 48.8 ⫾16.5 Glycolysis and TCA cycle enzymes Pfk NM ⬍0.05 ⬍0.05 Cs 32.5 ⫾22.3 71.7 ⫾38.7 66.5 ⫾40.9 Icdh 462 ⫾189 645 ⫾295 554 ⫾183 Anaplerotic pathway enzymes Icl 1.27 ⫾0.69 2.30 ⫾1.24 1.93 ⫾0.70 Mae 5.69 ⫾2.86 7.99 ⫾6.83 5.87 ⫾2.18 Pc 92.2 ⫾19.1 70.6 ⫾8.2 167.0 ⫾28.0 Ppc NM 9.02 ⫾3.67 15.82 ⫾4.42 Nitrogen metabolic enzymes NADPH-Gdh 22.3 ⫾7.60 73.7 ⫾31.4 16.4 ⫾9.0 NADH-Gdh 52.5 ⫾36.0 71.1 ⫾46.4 74.1 ⫾51.3 AspAT 95.2 ⫾18.0 174 ⫾93 78.5 ⫾16.5 Central Metabolism and Osmoadaptation in C. salexigens JUNE 14, 2013•VOLUME 288•NUMBER 24 JOURNAL OF BIOLOGICAL CHEMISTRY 17775 at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from The results obtained with [613 C]glucose, which were complementary to those obtained with [113 C]glucose, demonstrated that the incorporation of label into ectoines from C1 and C6 of glucose is not equivalent (supplemental Fig. S6). This result shows that glucose catabolism occurs through asymmetrically labeled 3-carbon metabolites, contrary to what would be predicted for the EM pathway. In addition, the more efficient incorporation of 13 C label into the glycerol moiety of membrane phospholipids from [613 C]glucose than from [113 C]glucose (supplemental Figs. S7 and S8) is also in accord with the labeling of ectoines. Thus, these data demonstrate that C. salexigens metabolizes glucose through the ED pathway and, together with the uncertainty of the existence of Pfk, suggest that the EM pathway is not functional. Ectoine Labeling in [213 C]Glucose-grown Cultures—For [213 C]glucose as carbon source, metabolism via the EM pathway would be predicted to yield PEP and pyruvate that are both 50% labeled in their C2. Metabolism via the ED pathway would also generate [213 C]pyruvate but would not produce any [ 13 C]PEP (supplemental Fig. S5). Therefore, the metabolic fate of the [213 C]pyruvate pool can be analyzed without any further assumptions (Fig. 5). OAA could be synthesized from pyruvate/ PEP in C. salexigens by the following routes: (i) carbons from pyruvate can enter the TCA cycle as acetyl-CoA produced by Pdh, rendering OAA after one turn of the cycle (Fig. 5A); (ii) pyruvate and PEP can be carboxylated to OAA by Pc or Ppc (Fig. 5B), or (iii) by a combination of both routes (Fig. 5C). From the spectra of ectoines, we can conclude that the contribution of Ppc to the total anaplerotic activity of the cells is negligible ( supplemental material). This would be in agreement with the measured enzyme activities (Table 3). Labeling of C6 and the carboxylic group of ectoines increase as a function of the Pdh flux, whereas labeling of C4 depends on the Pc flux. The labeling of C2 of ectoines is the result of the incorporation by EctA of the Pdh-produced acetyl-CoA. None of the pathway combinations would yield ectoines labeled at the methyl group. These predictions fit well with the corresponding spectra, where the signal coming from the methyl group is the least intense and the most intense signals were those of C2 and C4 (Fig. 5E). Effect of Salinity on Metabolic Fluxes, the Pc/Pdh, Cs/EctA, and Pc/Cs Flux Ratios To understand the functioning of central metabolic pathways in C. salexigens, the partitioning of pyruvate and acetylCoA can be described by the Pc/Pdh and Cs/EctA ratios. The Cs/EctA flux ratio describes the fraction of acetyl-CoA that enters the TCA cycle versus the fraction that is directly incorporated into ectoines. The Pc/Pdh flux ratio indicates the fraction of pyruvate that is transformed to OAA versus oxidized to acetyl-CoA. Finally, the Pc/Cs ratio allows comparison of the activity of anaplerosis and the TCA cycle and can be considered as readout of the biosynthetic or energetic needs of the cells (Fig. 3). The 13 C-labeling pattern of ectoines synthesized from [213 C]glucose can be used to quantify relative fluxes at these nodes (Table 4). For that aim, the metabolic steady state hypothesis was considered applicable to exponential cultures (and therefore labeling at the specific positions of the ectoine molecule is proportional to fluxes). Peak areas in 13 C NMR spectra were used to estimate flux ratios, using simple algebraic equations (supplemental material). The 13 C NMR spectra of FIGURE 4. 13 C NMR spectra of intracellular extracts of [113 C]glucose-grown cultures. M63 minimal medium with 0.75 MNaCl (upper spectrum) and 2.5 M NaCl (lower spectrum) was used. The signals corresponding to labeled carboxylic carbon (177 ppm for ectoine and 174–175 for hydroxyectoine) and C6 (38–39 ppm for ectoine and 44 for hydroxyectoine) are shown. In the scheme, the expected fate of labeled carbon when [113 C]glucose is metabolized via the Entner-Doudoroff pathway is shown. If we assume that the labeled C1 from pyruvate is lost as 13 CO 2 by decarboxylation at the level of Pdh and incorporated into OAA by Pc, the predicted ectoines labeling pattern would fit the spectra obtained. Relative labeling for each carbon atom is indicated by the color scale at right. The abbreviations used are as follows: labeled compounds detected: G, glutamate; E, ectoine; H, hydroxyectoine; GLC, glucose; PYR, pyruvate; OAA, oxaloacetate; AcCoA, acetyl-coenzyme A. Central Metabolism and Osmoadaptation in C. salexigens 17776 JOURNAL OF BIOLOGICAL CHEMISTRY VOLUME 288•NUMBER 24•JUNE 14, 2013 at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from FIGURE 5. Incorporation of label from [213 C]glucose into ectoines. 13 C from [213 C]pyruvate (derived via either the Embden-Meyerhof or the EntnerDoudoroff pathways), from [213 C]phosphoenolpyruvate (made via Embden-Meyerhof), or from unlabeled phosphoenolpyruvate (made via Entner-Doudoroff) can be incorporated into ectoines through the following. A, oxaloacetate synthesized in a single TCA cycle turn; B, oxaloacetate synthesized by pyruvate carboxylase or phosphoenolpyruvate carboxylase (Pc/Ppc), or (C) oxaloacetate synthesized by Pc/Ppc followed by a turn through the TCA cycle, which alters its labeling pattern (see the text for details). D, scheme depicting the relation of the pyruvate and acetyl-CoA nodes with the ectoines biosynthesis route in C. salexigens.E, 13 C NMR spectra from intracellular extracts of [213 C]glucose-grown cultures. M63 minimal medium with 0.75 MNaCl (upper spectrum) and 2.5 M NaCl (lower spectrum) was used. The signals corresponding to labeled carboxyl carbon, methyl carbon, C2, C4, and C6 of ectoine (E) and hydroxyectoine (H) are shown. The signal corresponding to hydroxyectoine carboxyl carbon overlaps with that of C1 of glutamate (indicated as C1, G). Note that for each pair of chemical shifts corresponding to each carbon, the ratio of the ectoine/hydroxyectoine signals is approximately constant. The three ectoine molecules in the inset represent the isotopomer distributions corresponding to a–c. Relative labeling for each carbon atom is indicated by the color scale at right of E. Where applicable, the upper half of the corresponding carbon position ball depicts the expected labeling from [213 C]pyruvate/[213 C]phosphoenolpyruvate, and the lower half from unlabeled phosphoenolpyruvate. See supplemental Table S4 and Fig. S5 and supplemental material “Determination of Metabolic Flux Ratios” for details. Central Metabolism and Osmoadaptation in C. salexigens JUNE 14, 2013•VOLUME 288•NUMBER 24 JOURNAL OF BIOLOGICAL CHEMISTRY 17777 at FAC BIOLOGIA/BIBLIOTECA on June 20, 2016http://www.jbc.org/Downloaded from