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Applied and Environmental Microbiology, April 2002, p. 1760-1771, Vol. 68, No. 4
0099-2240/02/$04.00+0     DOI: 10.1128/AEM.68.4.1760-1771.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.

Intracellular Carbon Fluxes in Riboflavin-Producing Bacillus subtilis during Growth on Two-Carbon Substrate Mixtures

Michael Dauner,1 Marco Sonderegger,1 Michel Hochuli,2 Thomas Szyperski,2,{dagger} Kurt Wüthrich,2 Hans-Peter Hohmann,3 Uwe Sauer,1* and James E. Bailey1

Institute of Biotechnology,1 Institute of Molecular Biology and Biophysics, ETH Zürich, CH-8093 Zürich, Switzerland,2 Roche Vitamins Inc., Nutley, New Jersey 07110-11993

Received 24 July 2001/ Accepted 8 January 2002


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ABSTRACT
 
Metabolic responses to cofeeding of different carbon substrates in carbon-limited chemostat cultures were investigated with riboflavin-producing Bacillus subtilis. Relative to the carbon content (or energy content) of the substrates, the biomass yield was lower in all cofeeding experiments than with glucose alone. The riboflavin yield, in contrast, was significantly increased in the acetoin- and gluconate-cofed cultures. In these two scenarios, unusually high intracellular ATP-to-ADP ratios correlated with improved riboflavin yields. Nuclear magnetic resonance spectra recorded with amino acids obtained from biosynthetically directed fractional 13C labeling experiments were used in an isotope isomer balancing framework to estimate intracellular carbon fluxes. The glycolysis-to-pentose phosphate (PP) pathway split ratio was almost invariant at about 80% in all experiments, a result that was particularly surprising for the cosubstrate gluconate, which feeds directly into the PP pathway. The in vivo activities of the tricarboxylic acid cycle, in contrast, varied more than twofold. The malic enzyme was active with acetate, gluconate, or acetoin cofeeding but not with citrate cofeeding or with glucose alone. The in vivo activity of the gluconeogenic phosphoenolpyruvate carboxykinase was found to be relatively high in all experiments, with the sole exception of the gluconate-cofed culture.


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INTRODUCTION
 
During batch growth on mixtures of carbon substrates, bacteria frequently first consume almost exclusively their preferred substrate. Consumption of any other substrate(s) occurs only after depletion of the preferred one, leading to a diauxic pattern of growth. The primary molecular mechanisms that inhibit the simultaneous utilization of substrates are catabolite repression (9) and inducer exclusion (47). Under carbon-limited conditions in chemostat cultures, in contrast, mixtures of carbon sources are often utilized simultaneously at low and intermediate dilution rates (D) (18). Thus, for the most frequently used paradigm catabolite repression sugar, glucose, such coutilization occurs at concentrations below a critical repression level (29). Consequently, many catabolic enzymes are relieved from repression so that alternative substrates can be catabolized (32, 34).

While many biotechnological processes operate on a single carbon source, substrate mixtures may help to diagnose potential bottlenecks in the biosynthetic pathways to a desired product (10). For the production of riboflavin (vitamin B2), which is a commercially important additive in the feed and food industries, such cofeeding experiments were used to identify potential limitations in building block supply (55) (Fig. 1). Generally, cofeeding experiments are evaluated on the basis of physiological analysis, with a primary focus on production. Much information on the underlying metabolic network response, however, is not accessible from extracellular physiological data but requires knowledge of intracellular carbon fluxes.



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  		FIG. 1. Purine and riboflavin biosynthesis pathway. Produced or consumed cofactors or building blocks are shown below the pathway. Abbreviations: P5P, pentose-5-phosphate; PRPP, phosphoribosylpyrophosphate; ~P, number of equivalents.

The classical approach to analyzing intracellular carbon fluxes is based on so-called metabolite balancing, which usually requires assumptions about redox or energy balances that strongly affect the flux estimates (8, 62). With the development of appropriate labeling experiments as well as extensive isotope isomer (isotopomer) models (11, 53, 63), a powerful tool for flux analysis was introduced that allows researchers to avoid such assumptions about redox and energy balances and thus increases the resolution and reliability of the estimates. The term "isotopomer" describes the different positional combinations of 12C and 13C atoms within a single molecule. The metabolic flux state determines the distribution of isotopomers in the intracellular metabolites which, in turn, determines the distribution of isotopomers in the amino acids (Fig. 2). Using nuclear magnetic resonance (NMR) or mass spectroscopy, subsets of these isotopomer pools may then be analyzed, for example, in the amino acids of hydrolyzed proteins (12, 58). Balancing the isotopomer pools of all intermediates in a biochemical reaction network enables an accurate mathematical description of the label distribution within the network. Combined with data on extracellular rates of substrate consumption and product formation, such rigorous accounting of 13C label distribution in the entire metabolic system provides novel information on intracellular reaction rates (fluxes) (Fig. 2).



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  		FIG. 2. The metabolic flux state is one determinant of the isotopomer pools in metabolic intermediates and amino acids. Additionally, the isotopomer pools are influenced by the factors shown in gray. By use of isotopomer balancing, the metabolic flux state may be estimated (broken arrows) from NMR- or mass spectroscopy-based isotopomer data and knowledge of the additional influential factors (shown in gray).

The aim of this study was to investigate metabolic flux responses to cofeeding of different carbon substrates in chemostat cultures of a recombinant, riboflavin-producing Bacillus subtilis strain (13, 41). For this purpose, four cosubstrates were chosen that enter central carbon metabolism at different positions. Gluconate and citrate provide direct intermediates of the pentose phosphate (PP) pathway and the tricarboxylic acid (TCA) cycle, respectively. Hence, they may serve as a carbon and/or energy source for riboflavin biosynthesis. Acetoin and acetate, in contrast, are catabolized via acetyl coenzyme A (acetyl-CoA) and, due to a lack of the glyoxylate shunt in B. subtilis (30), can serve only as an energy source by catabolism in the TCA cycle. To elucidate the intracellular flux responses to two-substrate catabolism, we performed biosynthetically directed 13C labeling experiments by growing chemostat cultures on mixtures of 90% unlabeled glucose, 10% [U-13C6]glucose, and one of the above unlabeled cosubstrates. The labeling patterns were assessed by two-dimensional (2D) NMR analysis of amino acids obtained by hydrolysis of cellular proteins (58). In contrast to previous analyses based on 2D NMR data (50, 51), we use here a recently developed comprehensive isotopomer model for data interpretation that enables more rigorous accounting of all labeling data (11, 14, 19).


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MATERIALS AND METHODS
 
Bacterial strain.
The recombinant, riboflavin-producing B. subtilis strain RB50::pRF69 was used throughout this study. The host strain, RB50 (purA60 Azr-11 Dcr-15 MSr-46 RoFr-50 spo0A), is characterized by several randomly introduced purine and riboflavin analog resistance mutations (41). To construct RB50::pRF69, one copy of the constitutively expressed, recombinant B. subtilis rib operon, pRF69, was integrated in the native chromosomal rib operon (41). To increase gene dosage, the pRF69 operon was amplified by selection in serial batch cultures up to a concentration of 80 mg of chloramphenicol liter-1. The resulting strain was denoted RB50::[pRF69]n, where n is the number of amplified pRF69 rib operons. While the exact number of amplifications is not known, all results presented are directly comparable because we inoculated all cultures from the same frozen stock of RB50::[pRF69]n, which was also used in previously reported experiments (11, 13, 14).

Growth conditions and media.
All media were supplemented with 80 mg of chloramphenicol liter-1. Frozen stocks were used to inoculate a complex medium, which contained 5 g of glucose liter-1, 25 g of veal infusion broth liter-1, and 5 g of yeast extract liter-1. After growth for 12 h in the complex medium, 2.5 ml was used to inoculate 50 ml of minimal medium in a 500-ml baffled shake flask; this mixture was cultivated for another 12 h and used entirely for reactor inoculation. The minimal medium used for the batch culture was similar to that used for the chemostat culture but did not contain H2SO4 and was supplemented with 0.1 M sodium phosphate buffer (pH 6.8).

Continuous cultivation in aerobic, glucose-limited chemostat cultures was conducted at 38°C with a working volume of 1 liter in a 2-liter LH discovery 210 series reactor (Adaptive Biosystems) equipped with pH, dissolved oxygen, temperature, optical density, and foam probes. The medium of the glucose-limited chemostat cultures contained (per liter of bidistilled water) the following: glucose, 5 g; NH4Cl, 4.75 g; (NH4)2SO4, 6 g; Na2HPO4 · 12H2O, 0.99 g; KH2PO4, 4 g; MgSO4 · 7H2O, 0.42 g; CaCl2 · H2O, 46 mg; FeSO4 · 7H2O, 9 mg; and 40 ml of a trace element solution. The trace element solution had the following composition (per liter of bidistilled water): MnCl2 · 4H2O, 2.25 g; ZnCl2, 1.32 g; CuCl2 · 2H2O, 0.34 g; CoCl2 · 6H2O, 0.5 g; Na2MoO4 · 2H2O, 0.5 g; and AlCl3 · 6H2O, 1.25 g. The medium was acidified to a pH of between 2 and 3 by the addition of H2SO4 (95 to 97%) and was sterilized by passage through a 0.2-µm-pore-size filter. For cofeeding experiments, 2 g of acetoin, gluconate, acetate, or citrate liter-1 was added. During cultivation, the pH was controlled at 6.8 ± 0.1, and the volume was kept constant by a weight-controlled pump. A constant airflow of 1 liter min-1 was achieved by a mass flow meter, and the agitation speed was set to between 600 and 1,000 rpm, ensuring dissolved oxygen levels above 40% under all conditions. All reported chemostat experiments were performed at the physiological steady state, which was defined as at least five volume changes under the same conditions and stable optical density and exhaust gas readings for at least three volume changes.

During the labeling experiments, the feed medium was replaced by an identical medium but with 10% (wt/wt) of the total glucose present as [U-13C6]glucose (13C, >98%; Isotech, Miamisburg, Ohio). Biomass aliquots for NMR analysis were withdrawn after 0.8 volume change, so that according to first-order washout kinetics, 45% of the biomass was fractionally labeled.

Analytical techniques.
Cell dry weight was determined from at least eight parallel 10-ml cell suspensions that were harvested by centrifugation, washed with distilled water, and dried at 80°C for 24 h to a constant weight. Protein, RNA, and glycogen contents of the cells were determined by colorimetric and enzymatic assays as described elsewhere (13). Concentrations of carbon dioxide and oxygen in the bioreactor feed medium and effluent gas were determined with a mass spectrometer (Prima 600; Fisons Instruments).

Extracellular glucose, succinate, pyruvate (PYR), and phosphoenolpyruvate (PEP) concentrations were determined enzymatically with commercial kits (Beckman) or as described by Bergmeyer (5). Acetate, acetoin, and 2,3-butanediol concentrations were determined by gas chromatography (5890E; Hewlett-Packard) with a Carbowax MD-10 column (Macherey-Nagel) and with butyrate as the internal standard. Organic acid, acetoin, and diacyl concentrations in culture supernatants were determined by high-pressure liquid chromatography with a Supelcogel C8 column (4.6 by 250 mm) (Sigma) and a diode array detector (Perkin-Elmer). Phosphoric acid (0.2 N) was used as the mobile phase at a flow rate of 0.3 ml min-1 and 30°C. Hexosamine concentrations in supernatants originating from both peptidoglycan and teichoic acid were determined by a colorimetric assay (24) with glucosamine for calibration. Riboflavin concentrations were determined by measuring the absorption at 444 nm (A444) in cell-free culture broth.

Intracellular concentrations of ATP and ADP were determined as described previously (60) with an ATP bioluminescence kit (HS II; Boehringer Mannheim). Briefly, a 10-ml cell suspension was rapidly (within 0.7 s) withdrawn with a syringe containing precooled glass beads (-20°C). Aliquots (40 µl) were transferred to an Eppendorf cup and immediately quenched by mixing with 160 µl of dimethyl sulfoxide. After the addition of 800 µl of ice-cold 25 mM HEPES buffer (pH 7.75), the samples were stored at -80°C and analyzed within 1 week. ATP concentrations were measured directly with a spectrofluorophotometer (RF-5001PC; Shimadzu) by detecting the light emission at 562 nm. ADP was converted to ATP by adding PYR kinase and PEP to 1,250 U/ml and 1 mM, respectively. After incubation for 30 min at 37°C, the resulting total ATP level was determined, yielding the ADP level as the difference between the two values. Standard solutions of ATP-ADP mixtures at a molar ratio of 8:1 in the range of 2 to 800 nM were used for calibration.

Description of the biochemical reaction network.
The previously described comprehensive isotopomer model of B. subtilis central carbon metabolism (11), with an H+-to-ATP ratio of 4 (20, 48), was extended to accommodate the situations arising with cofeeding of multiple alternative carbon sources. Specifically, gluconate is assumed to enter the central carbon metabolism via the consecutive reactions of gluconate permease (GntP), gluconate kinase (GntK), and 6-phosphogluconate dehydrogenase (GntZ) (21). Since B. subtilis lacks the Entner-Doudouroff pathway, it is assumed that gluconate is further metabolized through the PP pathway (40). The additional catabolic genes are organized in the gntRKPZ operon, which is transcribed from a gluconate-inducible promoter upstream of gntR. Expression is controlled by the regulator GntR, which is inactivated by gluconate (22). Like that in Escherichia coli, gluconate uptake in B. subtilis occurs via a proton symport mechanism (57).

Under glucose-limited conditions, acetyl-CoA synthetase is responsible for acetate utilization (27). In a single-step reaction, acetate is converted to acetyl-CoA at the expense of one equivalent of ATP, which is cleaved into AMP and PPi. Due to the large negative reaction enthalpy under physiological conditions ({Delta}G°', -88.2 kJ/mol), this reaction is assumed to be irreversible. Exchange fluxes, however, are considered to occur via phosphotransacetylase and acetate kinase, which usually catalyze acetate formation (54). The uptake of acetate most likely occurs via passive diffusion and/or a monocarboxylate transporter (28). Because acetate either diffuses across the cytoplasmic membrane in its protonated form or is taken up by an H+ symporter (28), we assume that one equivalent of H+ is transported per transported molecule of acetate.

Generally, acetoin degradation is assumed to be mediated by the 2,3-butanediol cycle (26). However, there is also experimental evidence that acetoin can undergo direct oxidative cleavage (33); hence, the existence of two alternative enzymatic routes, encoded by the acu and the aco gene clusters, was proposed (27). Later evidence indicated that the aco gene cluster, which encodes the acetoin dehydrogenase enzyme system, is the primary pathway for acetoin utilization (31). This pathway involves the cleavage of acetoin into acetate and acetaldehyde, with the simultaneous reduction of an acceptor molecule. In the next step, which is catalyzed by aldehyde dehydrogenase, the acetaldehyde generated is converted to acetate, with the reduction of one NADH. The two resulting acetate molecules are then introduced into the central carbon metabolism by acetyl-CoA synthetase. In our model, NAD+ is considered the electron acceptor for both reduction steps. Acetoin transport is assumed to occur by either passive or facilitated diffusion; thus, no metabolic energy expenditures are considered.

Citrate enters directly into the TCA cycle. In an aconitase mutant of B. subtilis, the intracellular accumulation of citrate was reported to lead to the induction of a divalent cation-dependent citrate transport system (64). This transport system was described later as a citrate-Mg2+ symporter (6). Because transport depends on the delta pH component of the proton motive force, the number of symported protons was assumed to be 2 (6).

NMR spectroscopy data analysis.
To prepare samples for NMR spectroscopy, a culture aliquot was harvested and centrifuged at 1,200 x g for 20 min at 4°C. The cell pellet was washed once with 20 mM Tris-HCl (pH 7.6), centrifuged again, resuspended in 6 ml of 6 M HCl, and hydrolyzed by incubation in sealed Pyrex glass tubes for 24 h at 110°C. The hydrolysate was filtered through a 0.2-µm-pore-size filter and lyophilized. The dried material was dissolved in 600 µl of 20 mM deuterochloric acid in 2H2O, incubated for 2 h at room temperature, centrifuged, and used to record the NMR spectra.

2D heteronuclear 13C-1H correlation NMR spectra ([13C,1H]COSY spectra) were recorded at 40°C and a 13C resonance frequency of 150.8 MHz by using a Bruker DRX500 spectrometer as described previously (58). For each experiment, [13C,1H]COSY spectra were recorded for the aliphatic 13C-1H moieties (data size, 4,096 by 512 complex points; t1max = 373 ms; t2max = 72 ms) and the aromatic 13C-1H moieties (data size, 1,536 by 512 complex points; t1max = 393 ms; t2max = 72 ms) (58), with measurement times of 4.5 to 6 h and 2.5 h per spectrum, respectively. As described previously, the relative intensities of the different 13C-13C scalar coupling fine structures in the [13C,1H]COSY spectra were evaluated by integration with the program FCAL, version 2.3.0 (59).

Estimation of intracellular carbon fluxes.
For the quantification of carbon fluxes in central metabolism, the relative abundances of the 13C-13C scalar coupling fine structures corresponding to the individual amino acid carbon positions, the experimentally determined macromolecular biomass composition, and the physiological data were combined within an isotopomer balancing model. Precursor demands for biomass formation and the experimentally determined macromolecular composition were deduced from a previously published growth model (13). Briefly, the isotopomer balances of all metabolites that are represented in the model are calculated starting from a randomly chosen flux distribution. Superposition of 13C-13C scalar coupling fine structures corresponding to this isotopomer distribution is then used for simulation and comparison to the experimentally determined NMR spectra. The quality of the fit is judged by the {chi}2 (error) criterion. Through an iterative process of flux estimation and signal fitting, a flux solution is sought that corresponds to a minimal {chi}2 value. This optimal solution represents the maximum-likelihood flux distribution in the investigated metabolic system that reflects both the physiological data and the 2D NMR data.

To identify the global error minimum in the solution space by this iterative procedure, a range-restricted evolutionary algorithm was used (3). The final solution was obtained by restarting a modified direction set search algorithm according to Powell's quadratic convergent method at the optimal flux solution identified by the global search evolutionary algorithm (44). The parameter search was initiated from at least five different starting points, which always yielded similar results, with fluxes varying maximally ±10%. The solution with the lowest {chi}2 value was then subjected to detailed statistical analysis. For this purpose, the Jacobian matrix of the overall output function, i.e., the linear approximation of the nonlinear isotopomer balancing model, was calculated numerically at the flux solution obtained. Conclusions could then be drawn about how the measured state variables would be influenced by differential changes in the flux estimates. After the model was linearized, linear statistical theory could be applied (39) to calculate confidence intervals for single parameters (11).

Determination of the biochemical energy content.
To determine the biochemical energy content of carbon substrates, the steady-state flux balance was formulated as an optimization problem in which the production of a particular product or quotients of the production of pairs of products were maximized, subject to

(1)
where S is the stoichiometric matrix, {nu} is the vector of fluxes, and b is the rate vector of metabolite production. For this purpose, either a variant of Mehrotra's predictor-corrector algorithm (35), a primal-dual interior point method, or a sequential quadratic programming method was used, as implemented in the linprog and fmincon functions of the MATLAB Optimization Toolbox (The MathWorks, Inc.).


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RESULTS
 
Physiological analysis.
Continuous cultures of B. subtilis RB50::[pRF69]n were grown in minimal medium at D of 0.10 to 0.12 h-1 with either glucose or mixtures of glucose and gluconate, acetoin, acetate, or citrate as the growth-limiting substrates. In all instances, cometabolism of the additional substrates with glucose was observed (Fig. 3). While gluconate and acetate were consumed completely, only a fraction of the supplemental acetoin and citrate was used, with residual concentrations of 16 mM (71% of the supplied acetoin) and 0.9 mM (8% of the supplied citrate) in the effluent medium, respectively. The specific substrate uptake rate in terms of carbon moles (C-mol) was enhanced in all cofeeding experiments compared to the single-substrate experiments with glucose at the same D (Fig. 3). This result was also reflected in the specific oxygen consumption and carbon dioxide evolution rates (data not shown). Generally, the formation of by-products accounted for 4 to 8% of metabolized total molar carbon, and no significant differences were observed between single-substrate and mixed-substrate cultures (data not shown).



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  		FIG. 3. Specific uptake rates for glucose, cofed substrates, and total carbon in carbon-limited chemostat cultures. Cultures were grown either on glucose only or on mixtures of glucose and the indicated cosubstrates.

As may be expected, the biomass yield on glucose was significantly higher in the cofeeding experiments because additional carbon sources were metabolized. The sole exception was a slightly reduced yield with acetoin as the cofed substrate. Specifically, the following yields (in grams mole-1) were obtained: 71 (acetate; D, 0.1 h-1), 64 (citrate; D, 0.1 h-1), 73 (gluconate; D, 0.12 h-1), and 55 (acetoin; D, 0.12 h-1); in comparison, yields were 56 and 57 with glucose only at D of 0.1 and 0.12 h-1, respectively. When normalized to the total carbon content in the substrates, however, the biomass yield was consistently lower in all instances (Fig. 4). To account for the different degrees of reduction (or energy) in the substrates as well, two different yield coefficients were calculated to attempt to normalize the biomass yield to the energy content of the substrate. The first yield coefficient, YX/e, is defined as the weight of dry cell mass produced per electron equivalents on n substrates i(Si) according to

(2)
where YX/Si is the molar yield coefficient on Si during the cofeeding experiment and Ye/Si is the number of electrons available from complete combustion of the substrate to CO2 (7). Ye/Si can be deduced from the degree of reduction ({gamma}S) (45) by multiplication with the number of carbon atoms contained in Si; the {gamma}S values are 4 for glucose, 3.67 for gluconate, 4 for acetate, 5 for acetoin, and 3 for citrate. The second yield coefficient, YX/BEC, is based on the same principle but considers the specific cellular biochemistry and is a more accurate means to differentiate energy-deficient and energy-excess substrates (1, 2), according to

(3)
where YBEC/Si is the biochemical energy content of Si.YBEC/Si was calculated as the maximum amount of ATP that could be generated from a substrate molecule within the biochemical reaction network of B. subtilis. On average, per carbon atom, a maximum of 2.50 molecules of ATP were found for glucose, 2.21 were found for gluconate, 1.13 were found for acetate, 1.75 were found for acetoin, and 1.58 were found for citrate. Consistent with the carbon-based yield, in almost all instances when the energy content was considered (YX/e and YX/BEC), the biomass yields were lower for the mixed-substrate cultures than for the cultures with glucose alone. Thus, growth on the mixtures was obviously less efficient than that on glucose alone.



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  		FIG. 4. Biomass yields and riboflavin yields in chemostat cultures. The biomass yields are calculated per mole of carbon (left, dark gray bars), mole of electrons (middle, light gray bars), or mole of ATP (right, black bars) available in the substrates.

While the biomass yield decreased in all cofeed scenarios, the yield of riboflavin showed no such consistent trend (Fig. 4). Cofeeding of acetoin or gluconate, respectively, resulted in 64 or 29% higher yields of riboflavin per mol of carbon in the consumed substrates. While citrate had no significant effect, the acetate-glucose mixture reduced the riboflavin yield by about 50%.

Metabolic flux analysis.
To gain deeper insight into the intracellular carbon flux distribution during dual-substrate limited growth, metabolic flux analysis was performed for each of the aforementioned experiments. First, to assess the building block requirements for biomass formation, we determined the contents of cellular protein, RNA, and glycogen. Glycogen was not detected, while the protein and RNA contents were virtually identical in all cultures, accounting for 50 to 55% and 6 to 8% of cellular dry weights, respectively (data not shown). On the basis of these data and a previously developed structured biomass model for chemostat growth of B. subtilis RB50::[pRF69]n (13), we calculated the biomass composition and the corresponding detailed building block requirements. Using the linear function of biomass composition with D from this structured model, which is based on 17 chemostat runs, we reduce potential inaccuracies from individual experimental analyses. In the single-substrate experiment, the carbon balance was 106.1% ± 4.3%, and in the cofeeding experiments with gluconate, acetoin, acetate, and citrate, the carbon balances were 98.2% ± 3.8%, 98.4% ± 3.4%, 103.6% ± 4.0%, and 107.9% ± 4.4%, respectively.

In the physiological steady state, all cultures were subjected to a labeling experiment during which the normal medium was replaced by a similar medium containing 10% (wt/wt) uniformly 13C-labeled glucose. After about 0.8 volume change, aliquots of cells were harvested and [13C,1H]COSY spectra were recorded for the total hydrolyzed biomass (58). The intracellular flux distribution was calculated as the best fit to the relative intensities of the 13C-13C scalar coupling fine structure multiplets for 44 carbon atoms in the amino acids (Table 1 and Appendix), detailed building block requirements, and extracellular fluxes. This procedure was carried out with a previously described, comprehensive isotopomer model of B. subtilis metabolism (11). The flux distribution in the two glucose-limited experiments at D of 0.10 and 0.12 h-1 were virtually identical, and in the following experiments, the latter was used as the reference experiment (Fig. 5A).


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  		TABLE 1. Carbon atoms subjected to 2D COSY NMR analysis in fractionally 13C-labeled amino acids and their corresponding precursors in central metabolism


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  		APPENDIX Measured and simulated values of the 2D COSY spectra of amino acidsa




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  		FIG. 5. Metabolic flux distribution of B. subtilis RB50::[pRF69]n in carbon-limited chemostat cultures with glucose (A), glucose and gluconate (B), glucose and acetate (C), glucose and acetoin (D), and glucose and citrate (E). D was 0.1 h-1 (C and E) or 0.12 h-1 (A, B, and D). Gray arrows indicate precursor withdrawal for biomass biosynthesis. Net and exchange fluxes (millimoles gram-1 hour-1) are given in square and oval boxes, respectively. Due to the fitting procedure used for intracellular flux estimation, extracellular fluxes can show small deviations from the experimentally determined data (Fig. 3). Abbreviations: G6P, glucose-6-phosphate; F6P, fructose-6-phosphate; 6PG, 6-phophogluconate; E4P, erythrose-4-phosphate; T3P, triose-3-phosphate; PGA, phosphoglycerate; ACA, acetyl-CoA; OGA, oxoglutarate; MAL, malate; FUM, fumarate; CIT, citrate; n.d., not determined.

As a measure of in vivo enzyme activity, the absolute fluxes (millimoles gram-1 hour-1) were found to vary significantly between the different experiments, most markedly for the PP pathway (glucose-6-phosphate to pentose-5-phosphate), PEP carboxykinase (oxaloacetate [OAA] to PEP), malic enzyme (malate to PYR), and the anaplerotic PYR carboxylase (PYR to OAA) (Fig. 5). The latter activity was particularly low with citrate cofeeding (Fig. 5E), presumably because citrate is the only substrate that can directly replenish TCA cycle intermediates; thus, the anaplerotic reaction is not required. Although citrate supply was more than sufficient to replenish the withdrawal of TCA cycle intermediates for building block biosynthesis, we found low but significant anaplerotic PYR carboxylase activity. Significant malic enzyme activity was seen only during cofeeding with gluconate, acetate, and acetoin. The gluconeogenic PEP carboxykinase was active in all experiments, except during gluconate cofeeding. Generally, the in vivo activity of the TCA cycle was higher in all cofeeding experiments than in the reference experiment, with the highest activity occurring during cofeeding with acetate or acetoin (Fig. 5C and D).

In some instances, it is informative to consider fluxes relative to the glucose uptake rate to allow a more direct comparison between experiments with different uptake rates. Although the absolute fluxes through the oxidative PP pathway were shown to vary considerably, from 0.10 to 0.42 mmol g-1 h-1 (Fig. 5A and C), the normalized fluxes were rather similar, ranging from 7% (acetate cofed) to 20% (glucose limited). Because gluconate is catabolized entirely via the oxidative PP pathway, it greatly increases the fluxes downstream of glucose-6-phosphate. The split ratios between the fluxes through the anaplerotic PYR carboxylase and the catabolic PYR dehydrogenase (PYR to acetyl-CoA) were found to span 1 order of magnitude over the five environmental conditions considered, from 1:1.7 to 1:17.5, illustrating the remarkable flexibility of the PYR branch point.

The obtained {chi}2 values, indicating the quality of the fit of the above flux estimates to the experimental data (Fig. 3 and 4 and Appendix), were 197 (acetoin), 624 (citrate), 646 (gluconate), and 481 (acetate); the value obtained with glucose as the sole carbon source was 119. The {chi}2 values in the citrate, gluconate, and acetate cofeeding experiments were relatively high, since typical values for glucose-grown B. subtilis or E. coli are about 100 when considered in the same type of labeling experiment and 2D COSY analysis (11, 19). Considering the degree of freedom within the single-substrate experiments, values of about 100 and 120 at confidence levels of 68 and 95%, respectively, typically would be expected (11). Closer inspection reveals that more than 50% of the total error criterion originates from deviation between simulated and measured 2D COSY data at very few carbon positions (Appendix). Specifically, these carbon positions are Tyr-{delta}x and {varepsilon}x for the citrate cofeed; His-ß, Lys-{gamma}, Ile-{gamma}1, Pro-{gamma}, Tyr-{varepsilon}x, and Glu-{gamma} for the gluconate cofeed; and six-His-ß, Arg-{delta}, Leu-ß, Lys-{varepsilon}, Tyr-{varepsilon}x, and Ile-{gamma}2 for the acetate cofeed. In almost all instances, poorly fitted signals originate from carbon positions that are located at the junction of different metabolic precursors for a particular amino acid. Obviously, NMR multiplet signals are very sensitive to slight variations in the degree of labeling in metabolic precursors; this situation occurs only in the cofeeding experiments. This problem is less apparent in the acetoin cofeeding experiment because the additional substrate is not used extensively for anabolism. This phenomenon was not considered in the present error model (11), so that the reported {chi}2 values are likely overestimated in the cofeeding experiments. Since the misfits vary between individual experiments, structural errors in the model are less likely than difficulties in automatic determination of multiplet NMR signals (59, 61). This view is also consistent with experimental signal differences between Pro-{gamma} and Glu-{gamma} and among Ile-{gamma}2, Val-{gamma}1, and Leu-{delta}1 in the gluconate and acetate cofeeding experiments, respectively (Appendix). These carbon positions originate from the same precursor metabolite; thus, one would expect similar signal ratios.

Bidirectional reaction steps and their impact on the labeling state of the system are included in the flux model, and the obtained estimates for the exchange fluxes are given in Fig. 5. For the experiments shown here, these estimates provide at best qualitative information, simply because variations in the exchange cannot be discerned from the available data. This fact has, however, only a marginal influence on the estimated net fluxes.

The reliability of the flux estimates was verified by two different procedures. First, the iterative fitting procedure was repeated at least five times with each data set. Although the parameter search was initiated from randomly chosen starting points, the resulting flux distributions were very similar in all instances (data not shown). Thus, there is a high probability that a global minimum was identified. Second, the linearized model was used to compute the confidence regions around the flux estimates of the reported solutions with the lowest {chi}2 values (11). The 68% confidence intervals of the oxidative PP pathway flux were ±20% the specific glucose uptake rate. Net fluxes in glycolysis (PGA to PEP) and the TCA cycle were more precisely determined, with 68% confidence intervals of ±10% and ±6%, respectively.

It is commonly observed that fluxes in the PEP-PYR-OAA triangle are difficult to resolve, and estimates typically show high confidence intervals (11, 43). As shown before, a linearized statistical model is not well suited to assessing the statistical relevance of these particular fluxes, since the confidence regions thus calculated are much larger than those obtained by other approaches (11). It appears, therefore, that the confidence intervals for the estimated fluxes in this triangle are on the order of magnitude of those for the other fluxes in the network. For more detailed analysis of these particular fluxes, it would be necessary to use labeled cosubstrates as well (43).

Redox and energy metabolism.
The estimated flux distribution also allows conclusions about intracellular redox and energy metabolism to be drawn. The majority of biosynthetic NADPH (26) was produced by the isocitrate dehydrogenase reaction in the TCA cycle. In the reference experiment and the gluconate cofeeding experiment, however, significant proportions of total NADPH were produced in the oxidative PP pathway (Fig. 6). On the demand side, the relative NADPH requirements for biomass formation and riboflavin biosynthesis are reduced in the cofeeding experiments (Fig. 6). Consequently, higher proportions of NADPH are converted to NADH by the transhydrogenase reaction, which was previously shown to be active in B. subtilis (13). This flux of reducing equivalents is a direct function of the estimated carbon fluxes; hence, it is determined within the confidence region of the estimated fluxes, in particular, the oxidative PP pathway flux. Although the 68% confidence interval of this particular flux is ±20% the specific glucose uptake rate, the data presented demonstrate clearly that a transhydrogenase or a transhydrogenase-like reaction must operate in B. subtilis metabolism.



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  		FIG. 6. Specific rates of NADPH production and consumption in carbon-limited chemostat cultures. NADPH production is accomplished via the oxidative PP pathway (lower, dark gray bars) and isocitrate dehydrogenase (upper, light gray bars). NADPH consumption occurs via biomass formation, transhydrogenase, and riboflavin biosynthesis (dark gray, light gray, and black bars, respectively).

Unlike NADPH production, metabolic production of ATP is not directly accessible from the flux distribution because the stoichiometry of ATP generation via respiration and ATPase is not known exactly and may vary with environmental conditions (49). It is not unreasonable, however, to assume identical efficiencies of ATP generation in all the carbon-limited samples investigated here, thus allowing for a direct comparison of ATP production in the five experiments. Specifically, we used a P-to-O ratio of about 1, which corresponds to the generation of 1 ATP molecule per NADH (13, 49). Depending on the estimated flux through the succinate dehydrogenase that produces the energetically less valuable FADH, however, the P-to-O ratio may be lower than 1. From the fluxes depicted in Fig. 5, one thus obtains the ATP production rate (Fig. 7). An almost invariant fraction of the produced ATP is required for the biosynthesis of biomass and riboflavin, and a comparatively minor fraction is required for futile cycle reactions, such as those of the PEP carboxykinase and the malic enzyme (Fig. 7). By far the largest fraction of ATP produced, however, is not required for any specific purpose and is thus referred to as excess ATP. The absolute excess production of ATP was slightly but significantly increased in all but one of the cofeeding experiments, with the acetate-cofed culture being the only exception. This result explains the higher YX/BEC with acetate and the lower YX/BEC in the other cofed cultures compared to the data in the reference experiment. The different behavior of the acetate-cofed culture is not related to a potentially lower P-to-O ratio, for example, through a decoupling of the proton gradient, because the residual acetate concentration was below the detection level and the proton-coupled uptake of acetate was already considered in the calculation.



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  		FIG. 7. Specific ATP production rates in carbon-limited chemostat cultures. The ATP fraction consumed for biomass (including riboflavin) formation and futile cycling via malic enzyme and PEP carboxykinase and the excess ATP fraction are indicated by light gray, black, and dark gray bars (top to bottom), respectively.

To investigate whether reduced growth efficiency and increased excess ATP production were correlated with the cellular energy state, we determined the intracellular ATP and ADP concentrations in the physiological steady state prior to the labeling experiment. All investigated cofeeding experiments were characterized by high ATP levels, which were more than doubled in the gluconate and acetoin cofeeding experiments compared to the reference experiment (Fig. 8). These two cultures exhibited unusually high ATP-to-ADP ratios as well.



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  		FIG. 8. Cellular contents of ATP (left, light gray bars) and ADP (right, dark gray bars) in carbon-limited chemostat cultures. CDW, cell dry weight.


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DISCUSSION
 
In this study, we used an isotopomer model to obtain a best-fit flux solution to all physiological data and the relative abundances of the 13C-13C scalar coupling fine structure multiplets for 44 amino acid carbon positions obtained from biosynthetically directed fractional 13C labeling experiments. The split ratio of glycolysis to PP pathway was relatively invariant at about 80% in all experiments for growth on one- and two-carbon substrate mixtures. Surprisingly, coutilization of gluconate did not reduce this split ratio significantly, although gluconate supply exceeded by far the cellular demands of NADPH and pentoses. Hence, it appears that flux into the PP pathway of B. subtilis is not controlled by the cellular demand for NADPH and/or pentoses but rather is determined by the kinetic properties of the enzymes at this branch point, as was previously shown for Corynebacterium glutamicum as well (36). The PYR node, in contrast, was rather flexible (56), because cofeeding with citrate reduced the anaplerotic flux via PYR carboxylase almost sixfold. The in vivo activity of the TCA cycle can respond in a flexible manner to cellular requirements. This fact is illustrated by more than twofold variations in the estimated TCA cycle flux on different substrate mixtures.

The gluconeogenic PEP carboxykinase was found to be active under all conditions investigated. Although the expression of this enzyme had been shown to be glucose repressed in B. subtilis batch cultures (16), very low residual glucose expression in our chemostat cultures was obviously not sufficient to exert complete repression. Consistent with this view, it was reported previously that in vivo PEP carboxykinase activity increased with decreasing glucose concentrations at lower growth rates (11, 50). Regulation of this in vivo activity, however, is more complex because ammonia- or phosphate-limited chemostat cultures exhibited intermediate or absent PEP carboxykinase fluxes, respectively, although high extracellular glucose concentrations were present in both cultures (14). Similar to in vitro results obtained with E. coli (37), we found significant in vivo malic enzyme activity on a glucose-acetate mixture. Likewise, high in vivo activities occurred with gluconate and acetoin as cosubstrates but not with citrate-glucose cofeeding or on glucose alone. From these results, we conclude that neither increased futile cycle activity via PEP carboxykinase or malic enzyme nor substantially altered pathway fluxes, for example, through altered splitting between glycolysis and the oxidative PP pathway, are primarily responsible for the reduced growth efficiency in the mixed-substrate cultivations.

In addition to carbon fluxes, intracellular concentrations of intermediates are an important component of comprehensive metabolic analyses. Of particular importance are the concentrations of ATP and ADP, which reflect the cellular energy state and are also common regulators in many cellular activities (26, 52). All cofed cultures exhibited significantly higher concentrations of ATP and higher ratios of ATP to ADP than the glucose-limited culture (Fig. 8). Thus, cofeeding of alternative substrates improves the energy charge of B. subtilis. The flux results indicate, however, that this apparent energetic prosperity is mostly used for excess ATP formation which, in turn, is used for maintenance metabolism and energy-spilling reactions (46) that cannot be assessed by the present methodology (Fig. 7). This view would be consistent with the reduced biomass yield in all cofeeding experiments. Yet another explanation could be that the actual P-to-O ratio in B. subtilis is lower than the value of about 1 used in our model, a situation which would lead to reduced excess ATP production.

Several studies on rRNA synthesis control have indicated that the levels of nucleoside triphosphate pools increase linearly with the growth rate and, in particular, that the ATP pool may be responsible for growth rate-dependent regulation of rRNA synthesis (4, 23, 38). Although this view has recently been questioned (42), all these studies relied on alternative carbon sources to achieve different growth rates in batch cultures. Our findings indicate that differences in cellular ATP and ADP contents may depend not only on the specific growth rate but also on the substrates and the associated flux distribution. However, we cannot exclude the possibility that the apparent divergence with our results is partly related to our use of an industrial B. subtilis strain with several deregulatory mutations in the purine biosynthesis pathway (41).

B. subtilis does not contain the enzymes of the glyoxylate shunt and therefore cannot grow on acetate or acetoin as a single carbon source (30). Hence, these cosubstrates are expected to influence primarily energy metabolism, while gluconate can serve as a precursor for riboflavin biosynthesis. Acetoin and gluconate cofeeding increased the riboflavin yield by 64 and 29%, respectively, but citrate had no significant effect, and acetate addition even reduced the riboflavin yield by about 50% (Fig. 4). In particular, the very different metabolic responses in the acetate and acetoin cofeeding experiments are surprising because both substrates enter metabolism at the same intermediate, acetyl-CoA. The low yield with the acetate cofeed cannot be due to toxic effects of acetate because the effective acetate concentration in the chemostat culture was close to zero. It is important to note in this context that the two beneficial cosubstrates increased the intracellular ATP-to-ADP ratio most strongly. Due to the phosphorylation steps involved in the conversion of GMP to GTP, a strong interdependence on the ATP-to-ADP ratio can be suspected. Thus, the high ATP-to-ADP ratio in the gluconate and acetoin cofeeding experiments may be the primary reason for the improved riboflavin yield.

Cofeeding experiments are very useful to redirect carbon flow, such that by-product formation can be reduced and important overflow reactions may be identified (10). For example, cometabolism of glucose and citrate in batch cultures and continuous B. subtilis cultures was shown to prevent the formation of overflow products and to increase the carbon yield more than twofold (25). Moreover, the simultaneous use of glucose and gluconate in batch cultures of a recombinant D-ribose-producing B. subtilis strain led to higher yields of D-ribose and by-products (15). Our interest, however, was to evaluate the influence of cofeeding on yields under process-relevant conditions. Hence, we conducted the experiments such that little or no by-product formation occurred. It is difficult to compare the efficiencies of biomass and product formation on mixtures of substrates because carbon and energy contents are usually different. For industrial purposes, the most appropriate method is the calculation of product and biomass yields on a cost basis. For the discovery of mismatches in the regulation of carbon flow, however, physiologically meaningful concepts are required. The optimal approach would be the determination of biomass and product yield coefficients for individual substrates and the evaluation of whether the mixtures have purely additive effects (17). However, many substrates, such as acetate and acetoin in the present investigation, do not support growth when used as the only carbon and energy source (30). Thus, in addition to calculating biomass yields on a per-mole-of-carbon basis, we used two alternative methods that consider the energy content as well, YX/e and YX/BEC. Generally, all calculated biomass yields for B. subtilis RB50::[pRF69]n were reduced in the cofeeding experiments, indicating less efficient utilization of the mixtures than of glucose alone (Fig. 4). From a bioenergetic perspective, glucose alone is clearly the most efficient substrate for biomass formation by B. subtilis. For producing riboflavin, however, cofeeding acetoin is a pertinent strategy to improve energetic prosperity and thus the production of the desired compound.


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ACKNOWLEDGMENTS
 
This work was supported by a scholarship from the Boehringer Ingelheim Fonds to M.D., by Roche Vitamins Inc., and by the Swiss Priority Program in Biotechnology (SPP BioTech).


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FOOTNOTES
 
* Corresponding author. Mailing address: Institute of Biotechnology, ETH Zürich, CH-8093 Zürich, Switzerland. Phone: 41-1-633 36 72. Fax: 41-1-633 10 51. E-mail: sauer{at}biotech.biol.ethz.ch. Back

{dagger} Present address: Department of Chemistry, State University of New York at Buffalo, Buffalo, NY 14260. Back


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Applied and Environmental Microbiology, April 2002, p. 1760-1771, Vol. 68, No. 4
0099-2240/02/$04.00+0     DOI: 10.1128/AEM.68.4.1760-1771.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.



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