The energetic efficiency of microbial growth is significantly
reduced in cultures growing under glucose excess compared to cultures
growing under glucose limitation, but the magnitude to which different
energy-dissipating processes contribute to the reduced efficiency is
currently not well understood. We introduce here a new concept for
balancing the total cellular energy flux that is based on the
conversion of energy and carbon fluxes into energy equivalents, and we
apply this concept to glucose-, ammonia-, and phosphate-limited
chemostat cultures of riboflavin-producing Bacillus
subtilis. Based on
[U-13C6]glucose-labeling experiments and
metabolic flux analysis, the total energy flux in slow-growing,
glucose-limited B. subtilis is almost exclusively
partitioned in maintenance metabolism and biomass formation. In
excess-glucose cultures, in contrast, uncoupling of anabolism and
catabolism is primarily achieved by overflow metabolism, while two
quantified futile enzyme cycles and metabolic shifts to energetically
less efficient pathways are negligible. In most cultures, about 20% of
the total energy flux could not be assigned to a particular
energy-consuming process and thus are probably dissipated by processes
such as ion leakage that are not being considered at present. In
contrast to glucose- or ammonia-limited cultures, metabolic flux
analysis revealed low tricarboxylic acid (TCA) cycle fluxes in
phosphate-limited B. subtilis, which is consistent with
CcpA-dependent catabolite repression of the cycle and/or
transcriptional activation of genes involved in overflow metabolism in
the presence of excess glucose. ATP-dependent control of in vivo
enzyme activity appears to be irrelevant for the observed differences
in TCA cycle fluxes.
 |
INTRODUCTION |
The very basis of microbial
growth resides in balanced fluxes through anabolic and catabolic
reactions. These metabolic fluxes are highly variable and change with
the environmental conditions and the rate of growth, since
faster-growing cells demand a higher rate of metabolism. To delineate
these influences, metabolic flux responses are typically studied in
chemostat cultures that are maintained under different nutrient
limitations. When microorganisms are limited for their energy source
(usually the carbon source), catabolism is tightly coupled to anabolism
and high biomass yields on the carbon source are achieved
(40). Compared to those with carbon (C) limitation,
excess-C cultures exhibit generally high rates of carbon consumption
and low yields of biomass and thus have a low energetic growth
efficiency (11, 31, 32, 57). Most frequently excess-C
cultures in chemostats are limited by other cellular macroelements such
as nitrogen (N) and phosphorus (P) but also potassium, the predominant
intracellular cation (11). Potassium or P limitation
invokes usually a strong uncoupling of catabolic and anabolic processes
that lead consequently to low biomass yields on the energy source,
while N limitation has more moderate effects on cellular physiology
(31, 68). Although mostly studied in the gram-negative
Enterobacteria, these findings are consistent with data from
gram-positive Bacillus spp. (4, 28, 32, 35).
The uncoupling of anabolism and catabolism in excess-C cultures reduces
the energetic growth efficiency of these cultures, which is attributed
to so-called overflow metabolism, metabolic shifts of carbon or
electron flow to less efficient pathways, and/or a variety of
energy-spilling reactions (40, 57). While the magnitude of
ATP dissipation via energy-spilling reactions is not well understood,
several such processes that catalyze net loss of energy though cyclic
reactions are known at the molecular level. Such cycles are frequently
referred to as futile, simply because no clear metabolic function of
such apparent waste of energy could be envisaged (22, 30,
40). These futile cycles may result from ion or proton leakage
across the cytoplasmic membrane that reduces the proton motive force
and thus decreases the efficiency of respiratory ATP generation.
Another type of futile cycling is based on metabolic reactions that act
in an antagonistic fashion, so that one molecule of ATP is dissipated
per cycled metabolite. While ion leakage may contribute significantly
to energy spilling (30, 40), ATP-dissipating futile enzyme
cycles are generally thought to be maintained at a relatively low level
(5, 6, 8). In certain biological systems, however, futile
enzyme cycling is extensive (22). The most prominent of
these is probably thermogenesis in bumblebees during cold weather. More
recently, metabolic flux analysis revealed that certain futile enzyme
cycles may account for ATP dissipation in the order of one molecule of
ATP per consumed molecule of glucose in microbial cultures
(37; M. Emmerling, M. Dauner. A. Ponti, J. Fiaux, M. Hochuli, T. Szyperski, K. Wüthrich, J. E. Bailey, and U. Sauer, submitted for publication.).
Most enzymes that are involved in potential futile cycles are subject
to complex allosteric control, so that their in vivo operation may be
assessed only by isotope-labeling experiments (22). Such
labeling experiments were traditionally conducted such that the
distribution of label was indicative of one particular futile cycle
(22, 59). A common problem of such analyses are corrections for alternative pathways and exchange reactions that may
also affect label distribution. The currently most advanced form of
metabolic flux analysis can quantitatively account for the use of
alternative pathways and exchange reactions on the distribution of
13C label throughout the whole metabolic system
of a cell (9, 50, 65). Hence, this methodology provides a
holistic view on cellular metabolism and not only quantifies metabolic
shifts and certain futile cycles but can also shed some light on the energetic aspects of growth. This is achieved by the use of
comprehensive isotope isomer (isotopomer) models that encompass all
relevant metabolic intermediates and appropriate computational tools
for data analysis (9, 63, 64).
The primary focus of this work was to quantify the magnitude of energy
dissipation via overflow metabolism, metabolic shifts, and futile
enzyme cycling in C-limited and excess-C Bacillus subtilis chemostat cultures, by using isotopomer-balancing (9) and
[U-13C6]glucose-labeling
experiments (55). Thus, for the first time we report here
intracellular carbon flux distributions in N- and P-limited B. subtilis.
| |
MATERIALS AND METHODS |
Strain.
Throughout this study, the recombinant,
riboflavin-producing B. subtilis strain
RB50::pRF69 was used. The host strain, RB50 (purA60 Azr-11
Dcr-15 MSr-46
RoFr-50 spo0A), contains several
chemically introduced purine and riboflavin analog-resistant mutations
(36). In RB50::pRF69, one copy of the
constitutively expressed, recombinant B. subtilis rib operon
pRF69 with the cat marker, was integrated in the native chromosomal rib operon. To increase gene dosage, the
recombinant rib operon was amplified by chloramphenicol
selection of RB50::pRF69 in serial batch cultures up to a
concentration of 80 mg/liter. The resulting strain is designated
RB50::(pRF69)n, where n
refers to the number of amplified pRF69 rib operons.
All cultures were inoculated from the same frozen stock. This
amplification has no impact on the present study other than diverting 1 to 3% of substrate carbon into riboflavin.
Growth conditions and media.
All media were supplemented
with 80 mg of chloramphenicol per liter. Frozen stocks were revived for
12 h in complex medium, containing (per liter) 5 g of
glucose, 25 g of veal infusion broth, and 5 g of yeast
extract. A 2.5-ml amount of this culture was transferred into 50 ml of
minimal medium in a 500-ml baffled shake flask. After another 12 h, the whole culture was used for reactor inoculation. The minimal
batch medium was similar to the chemostat medium but did not contain
H2SO4 and was supplemented
with 0.1 M sodium phosphate buffer (pH 6.8).
Chemostat cultures were grown at 38°C in 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 N-limited medium contained (per
liter) 8.5 g of glucose, 0.53 g of
NH4Cl, 0.67 g of
(NH4)2SO4,
2.54 g of Na2HPO4
· 12 H2O, 20.72 g of
KH2PO4, 0.42 g of
MgSO4 · 7 H2O, 46 mg
of CaCl2 · H2O, 9 mg
of FeSO4 · 7 H2O,
and 40 ml of a trace element solution containing (per liter) 2.25 g of MnCl2 · 4 H2O, 1.32 g of ZnCl2, 0.34 g of
CuCl2 · 2 H2O,
0.5 g of CoCl2 · 6 H2O, 0.5 g of
Na2MoO4 · 2 H2O, and 1.25 g of
AlCl3 · 6 H2O. To
achieve P limitation at the low or high dilution rate (D),
phosphate salts were added at the following concentrations (per liter):
0.00192 g of Na2HPO4
· 12 H2O and 0.01536 g of
KH2PO4 or 0.032 g of
Na2HPO4 · 2 H2O and 0.032 g of
KH2PO4, respectively.
Chloride salts were newly added at the following concentrations to
ensure identical ionic strength in all experiments (per liter):
1.9 g of KCl and 0.45 g of NaCl or 1.68 g of KCl and
0.3 g of NaCl for high or low D, respectively.
For labeling experiments, the medium was replaced by an otherwise
identical medium that contained the same glucose concentration as a
mixture of 90% (wt/wt) natural glucose and 10% (wt/wt)
[U-13C6]glucose
(13C, >98%; Isotech, Miamisburg, Ohio).
Biomass aliquots for nuclear magnetic resonance (NMR) analysis were
withdrawn after 0.8 volume changes, so that 45% of the biomass was
fractionally 13C labeled according to first-order
washout kinetics.
Chemostat media were acidified to pH 2 to 3 by addition of
H2SO4 (95 to 97%) and
sterilized by passage through a 0.2-µm-pore-size filter. During the
fermentation, the pH was maintained at 6.8 and the volume
was kept constant by a weight-controlled pump. A constant airflow of 1 liter/min was achieved by a mass flowmeter, and the agitation speed was
set to values between 600 and 1,000 rpm, ensuring dissolved oxygen
levels well above 30% in all cases. All data reported are from
cultures in physiological steady state, defined as at least five
culture volume changes under the same conditions and stable optical
density and off gas readings for at least three volume changes.
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. Cellular protein, RNA, and glycogen
content were determined as described elsewhere (10).
Glucose, succinate, pyruvate, and phosphoenolpyruvate concentrations
were determined enzymatically with commercial kits (Beckman) or as
described by Bergmeyer (3). Acetate, acetoin, and
2,3-butanediol concentrations were determined by gas chromatography
(5890E; Hewlett-Packard) on a Carbowax MD-10 column (Macherey-Nagel)
with butyrate as the internal standard. Additionally, organic acids,
acetoin, and diacyl were determined by high-performance liquid
chromatography on a Supelcogel C8 column (4.6 by 250 mm) (Sigma)
with a diode array detector (Perkin-Elmer). Phosphoric acid (0.2 N) was
used as mobile phase at a flow rate of 0.3 ml/min and 30°C.
Concentrations of carbon dioxide and oxygen in the bioreactor feed and
effluent gas were determined with a mass spectrometer (Prima 600;
Fisons Instruments).
Hexosamine concentrations in the supernatant, originating from both
peptidoglycan and teichoic acids, were determined colorimetrically (15), using glucosamine for calibration. Riboflavin
concentrations were determined as the absorption at 444 nm
(A444) in cell-free culture broth. The
carbon, hydrogen, and nitrogen composition of dried biomass was
determined with a CHN-elemental analyzer (EA1108; CE Instruments).
Prior to analysis, cell pellets were washed with bidistilled
H2O, dried in a freeze-dryer for 12 h, and
stored over silica gel (Sigma) for another 12 h in an evacuated desiccator.
Intracellular concentrations of ATP and ADP were determined as
described previously (60), using an ATP bioluminescence
kit (HS II; Boehringer Mannheim). For this purpose, 10 ml of culture broth was rapidly (within 0.7 s) withdrawn with a syringe
containing precooled glass beads (
20°C). Aliquots of 40 µl were
transferred to a plastic cup and quenched by mixing with 160 µl of
dimethyl sulfoxide. After addition of 800 µl of ice-cold 25 mM HEPES
buffer (pH 7.75), samples were stored at 80°C and analyzed within 1 week.
Two-dimensional proton-detected heteronuclear single quantum
13C-1H correlation NMR
spectroscopy
([13C,1H]-COSY) and data
analysis were done exactly as described previously (48,
55) with amino acids obtained from hydrolyzed biomass that was
harvested after the 13C-labeling experiments
(9, 46, 48). For each sample, one spectrum was recorded
for the aliphatic 13C-1H
moieties and one spectrum was recorded for the aromatic
13C-1H groups. The relative
abundances of 13C-13C
scalar coupling multiplets in the
[13C,1H]-COSY cross peaks
were evaluated with the program FCAL (56; R. Glaser,
2.3.0. ed., 1999).
Biochemical reaction network.
To describe N and P
limitation, the previously described comprehensive isotopomer model of
B. subtilis central metabolism with an
H+-to-ATP ratio of 4 was extended
(10). Ammonium uptake in C- and P-limited (excess-N)
cultures was assumed to proceed via energy-independent diffusion. Under
N limitation, uptake was catalyzed by an
NH4+/K+
antiporter (41) and K+ was
replenished by a H+/K+
symporter (2). Intracellular ammonium assimilation was
accomplished by glutamine synthetase and glutamate synthase
(51). Phosphate uptake in C- and N-limited (excess-P)
cultures was assumed to proceed via an energy-independent, low-affinity
system. Uptake under P limitation was catalyzed by an ABC-type
transporter that required one ATP per transported phosphate
(41).
Building block requirements for biomass were calculated from
experimental data and a previously developed structured model, in which
the cell wall was assumed to consist of 45% peptidoglycan and 55%
teichoic acids (10). Under P limitation, however, B. subtilis strongly reduces the P content by replacing teichoic acid
with teichuronic acids (23), and thus an average chain length of 7.5 (34) with the repeating unit
(GlcA)2-(GalNAc)2 (26) was assumed for P-limited cultures.
Estimation of intracellular carbon fluxes.
Metabolic flux
analysis was based on three different data sets: (i) substrate uptake
and product formation rates; (ii) macromolecular biomass composition;
and (iii) relative abundances of the
13C-13C multiplets of 47 amino acid carbon positions in the
[13C,1H]-COSY spectra.
The requirement of metabolic precursor for biomass formation was
deduced from the experimentally determined macromolecular composition
and a previously published growth model (10).
Intracellular carbon fluxes were then calculated as a best fit to the
three data sets within an isotopomer model (9). This model
enables rigorous accounting of the isotopomer pools through 25 metabolite equations, 502 isotopomer equations for metabolic intermediates, and 3,628 isotopomer equations for amino acids that
balance all positional combinations of 12C and
13C in the considered compounds. Exchange fluxes
via reversible reactions were quantitatively considered in the model.
In the first step, the isotopomer balances of all metabolites (Fig.
1) were calculated from a randomly chosen
flux distribution. Relative abundances of the
13C-13C multiplets in the
amino acids are then simulated from this isotopomer distribution and
compared to the determined multiplets. The quality of the fit to the
multiplets and the physiological data is judged by the
2-error criterion. Through an iterative
process of flux estimation and signal fitting, a flux solution that
corresponds to the minimal 2 value is sought.
This optimal solution represents the maximum-likelihood flux
distribution in the investigated metabolic system that reflects both
the physiological data and the
[13C,1H]-COSY data.
A range-restricted evolutionary algorithm was used to identify the
global error minimum by this iterative procedure (1). The
final solution was obtained by restarting a modified direction-set search algorithm at the optimal flux solution that was identified by
the evolutionary algorithm (38).
ATP balance.
Like the commonly used carbon balance, energy
balances can be constructed for cellular processes that account also
for the incorporation of energy source into cell material. The
metabolic energy equivalent of glucose is the maximum amount of ATP
that may be generated from complete oxidation of glucose to carbon dioxide. For aerobic organisms such as B. subtilis, this ATP
generation is dependent on the coupling efficiency of respiration and
ATP synthesis and is usually expressed as the so-called P-to-O (ATP per
oxygen) ratio. B. subtilis contains a three-branched
electron transport chain with different coupling efficiencies, but two of these branches are probably irrelevant in our well-aerated, glucose-grown cultures, i.e., the bd oxidase branch is
primarily active at low oxygen tension (67), and the
cytochrome c branch is inactive in glucose-grown cells
(27, 66). Hence, oxidation of NADH in our cultures is
probably catalyzed by the aa3 oxidase branch, which was also shown experimentally to be predominant during
vegetative growth of B. subtilis (25, 42).
Transport of one electron from NADH along this branch translocates two
protons (25, 61), and four protons are needed to generate
one ATP in the reaction catalyzed by ATP synthase (13,
43). Transport of one electron from FADH, however, translocates
only one proton (49). Consequently, all cultures were
assumed to exhibit a maximal P/O of unity (the exact value depends on
the amount of FADH that is produced in the tricarboxylic acid [TCA]
cycle), as was previously suggested for B. subtilis
(10, 44). This P-to-O ratio corresponds to the generation
of 1 ATP per NADH and 0.5 ATP per FADH, so that complete oxidation of
glucose to carbon dioxide yields 15 ATP. The total cellular energy flux
in ATP equivalents is therefore the specific glucose uptake rate
multiplied by 15.
Excretion of metabolic by-products such as acetate causes an energetic
loss to the cell because glucose cannot be oxidized completely to
carbon dioxide, and hence less ATP is generated from the available
glucose. To account quantitatively for the incorporation of glucose
into products and biomass, we calculated the ATP equivalent of each
product. First, the maximum ATP generation was calculated for complete
conversion of glucose to each considered product and carbon dioxide at
a P-to-O ratio of 1. The difference of this ATP amount to the 15 ATP
that are potentially available from complete oxidation of glucose
represents the ATP equivalents that are lost by formation of the
considered product. For example, complete conversion of one molecule of
glucose to two acetate and two carbon dioxide molecules yields 6.5 ATP
molecules at the assumed P/O (16). Thus, formation of one
molecule of acetate in glucose catabolism is equivalent to the loss of
4.25 ATP molecules. This calculation was done for each considered
product and accounts also for all ATP and reducing equivalents that are
generated in the conversion of glucose to each product.
| |
RESULTS |
Physiology of excess-C B. subtilis.
To
elucidate the efficiency of B. subtilis growth when abundant
carbon was available, we grew B. subtilis
RB50::(pRF69)n under N or P limitation at
dilution rates of 0.1 and 0.4 h
1 in chemostat
culture. These rates were chosen such that the low value was below the
critical D at which aerobic fermentation may occur
(19, 52) and the high value was well below the maximum growth rate of RB50::(pRF69)n in this
medium, 0.58 h1. Residual glucose
concentrations were 0.8 and 2.4 g/liter in N-limited culture and 7.2 and 1.8 g/liter in P-limited culture, at the low and high D,
respectively, thus illustrating successful establishment of excess-C
conditions. For C limitation we used previously reported
glucose-limited chemostat experiments with RB50::(pRF69)n from the exact same stock
culture (10).
Most significantly, the biomass yield on glucose was highest in the
C-limited culture and lowest in the P-limited culture, demonstrating
clearly the low energetic efficiency in the latter (Fig.
2). Consequently, the specific glucose
uptake rate was enhanced in excess-C cultures compared to C-limited
cultures and most pronounced under P limitation (Fig.
3). As described also for other organisms (24), increased respiration is one response to C excess
under N-limited conditions, illustrated here by higher specific rates of oxygen consumption and carbon dioxide formation (Fig. 3). Under P-limited conditions, in contrast, the rate of respiration is comparable to that under C-limited conditions. Thus, increased respiration does not appear to be a general response to C sufficiency in B. subtilis.
View larger version (34K):
[in this window]
[in a new window]
|
FIG. 2.
Biomass yields on glucose in C-, N-, or P-limited
chemostat culture of B. subtilis. The data for C-limited
cultures were taken from Dauner and Sauer (10).
|
|
View larger version (22K):
[in this window]
[in a new window]
|
FIG. 3.
Specific glucose (A) and oxygen (C) consumption rates as
well as acetate (B) and carbon dioxide (D) production rates of
B. subtilis during C-, N-, or P-limited chemostat
cultivation. The data for C-limited cultures were taken from Dauner and
Sauer (10).
|
|
Formation of low-molecular-weight by-products (excluding riboflavin) by
so-called overflow metabolism was low in C-limited culture
(10), moderate in N-limited culture, and extensive in P-limited culture (Table 1), as was
described for Bacillus spp. (28, 35) and
E. coli (31). The primary by-product was
acetate, but diacyl and acetoin formation was also significant. The
specific production rate of acetate was strongly influenced by the
growth rate and was highest under P limitation (Fig. 3). Not generally considered classical products of overflow metabolism, extracellular protein and cell wall components contributed 1 to 4% of the carbon balance (data not shown). Recombinant riboflavin production contributed very little to the carbon balance and showed no pronounced response to
the different environmental conditions (Table 1).
View this table:
[in this window]
[in a new window]
|
TABLE 1.
Relative contribution of metabolic (by)products to the
carbon balance of C-, N-, or P-limited B. subtilis chemostat
cultures at dilution rates of 0.1 and 0.4 h1
|
|
Biomass composition.
To obtain accurate information on biomass
precursor requirements for subsequent metabolic flux analysis, we
determined the relative contents of protein, RNA, and glycogen in our
cultures. While glycogen was negligible under all conditions, the
protein content was 55% ± 5% and 55% ± 3% in the N-limited
culture and 65% ± 8% and 60% ± 3% in the P-limited culture at low
and high D, respectively. The RNA contents of 6% ± 1% and
9% ± 1% in N-limited cultures and 4% ± 1% and 9% ± 1% in
P-limited cultures at low and high D values, respectively,
were low when compared to those in C-limited cultures
(10). The elemental biomass compositions that were
calculated from these macromolecular biomass data agreed well with data
from experimental CHN analysis (Table 2),
thus validating the results. The high protein content of the
slow-growing, P-limited culture correlates with its high nitrogen
content. Similarly, the low protein content of the fast-growing,
N-limited culture correlates with its low nitrogen content.
View this table:
[in this window]
[in a new window]
|
TABLE 2.
Comparison of the experimentally determined elemental
composition of B. subtilis biomass to the composition
calculated from the macromolecular biomass data in N- and P-limited
chemostat cultures
|
|
Metabolic flux analysis.
To quantify the intracellular carbon
flux distribution in excess-C cultures, labeling experiments with
[U-13C]glucose were performed with cultures in
physiological steady state. After about 0.8 volume changes, culture
aliquots were harvested and hydrolyzed biomass was analyzed by
[13C,1H]-COSY
(55). The intracellular flux distribution was then
calculated as the best fit to the relative abundances of
13C-13C coupling fine
structure multiplets of 47 amino acid carbon positions (data not
shown), the detailed building block requirements, and the extracellular
fluxes (Fig. 1). Using the previously described, comprehensive
isotopomer model of B. subtilis metabolism (9), we obtained very good fits to the data, which indicates that all data
sets are consistent with each other.
The calculated intracellular fluxes represent estimates for in vivo
enzyme activities. To facilitate direct comparison of flux
distributions under different conditions, we normalized the fluxes to
the glucose uptake rate of each culture (Fig. 3). Generally, the
intracellular flux distribution depends strongly on the choice of the
limiting nutrient and the growth rate; and four major shifts in
metabolic pathway usage were discernible (Fig.
4 and 5). First, the relative TCA cycle
flux was extremely low under P limitation at either D,
while the high TCA cycle flux in C- and
N-limited cultures was markedly increased at the low D
value. Second, a high relative malic enzyme flux of 27% was identified
in the fast-growing P-limited culture (Fig. 5). Combined with the low
TCA cycle flux under this condition, a flux distribution with reversed
flux from oxaloacetate (OAA) to malate (MAL) via the MAL dehydrogenase
was achieved, compared to the carbon flux usually encountered in
aerobic cultures. Third, the relative PP pathway flux was low in the
slow-growing N-limited culture, so that glucose catabolism proceeds
almost exclusively through the glycolytic pathway. The split ratio
between these two catabolic fluxes was otherwise relatively constant, with the possible exception of the slow-growing P-limited culture. Fourth, the gluconeogenic flux from OAA to phosphoenolpyruvate (PEP)
catalyzed by PEP carboxykinase (16) was entirely absent in
P-limited culture but was around 20 and 10% at low and high D, respectively, under C or N limitation.
View larger version (87K):
[in this window]
[in a new window]
|
FIG. 4.
Metabolic flux distribution in C-limited (top entry in
the boxes), N-limited (middle), or P-limited (bottom) chemostat culture
of B. subtilis at D = 0.1 h1. Fluxes are relative to the specific glucose
consumption rate of each culture shown in Fig. 3. Large arrowheads
indicate the primary flux direction, and small arrowheads indicate that
a reaction was considered reversible. Solid gray arrows indicate
withdrawal of building blocks for biomass formation. For C-, N-, and
P-limited cultures we recovered 107% ± 4%, 105% ± 5%, and 108% ± 6%, respectively, of the consumed carbon in the determined
products. Abbreviations: G6P, glucose-6-phosphate; F6P,
fructose-6-phosphate; P5P, pentose phosphates; E4P;
erythrose-4-phosphate; S7P, seduheptulose-7-phosphate; T3P,
triose-3-phosphate; PGA, 3-phosphoglycerate; SER, serine; GLY, glycine;
C1, methyl group bound to tetrahydrofolate; PEP, phosphoenolpyruvate;
PYR, pyruvate; ACoA, acetyl-Coenzyme A; OGA, 2-oxoglutarate; FUM;
fumarate; MAL, malate; OAA, oxaloacetate; and TCA cycle,
tricarboxylic acid cycle. The data for C-limited cultures were taken
from Dauner et al. (9).
|
|
View larger version (88K):
[in this window]
[in a new window]
|
FIG. 5.
Metabolic flux distribution in C-limited (top entry in
the boxes), N-limited (middle), or P-limited (bottom) chemostat culture
of B. subtilis at a D of 0.4 h1. Fluxes are relative to the specific glucose
consumption rate of each culture shown in Fig. 3. Large arrowheads
indicate the primary direction of flux in a given reaction, and small
arrowheads indicate that a reaction was considered reversible. Solid
gray arrows indicate withdrawal of building blocks for biomass
formation. For C-, N-, and P-limited cultures we recovered 97% ± 3%,
112% ± 7%, and 104% ± 7% %, respectively, of the consumed carbon
in the determined products. The data for C-limited cultures were taken
from Dauner et al. (9).
|
|
As a consequence of the above metabolic flux responses, two reaction
sequences at the glycolysis-TCA cycle interface conclude ATP-dissipating futile cycles or bypass reactions (Fig.
6). The first is a cyclic flux from OAA
via PEP and pyruvate (PYR) that promotes net loss of one ATP per turn
and is catalyzed by PEP carboxykinase, PYR kinase, and PYR carboxylase.
This futile enzyme cycle is active primarily in slow-growing N-limited
cultures (Fig. 4 and 5), as was previously described for C-limited
B. subtilis (9, 46). The second is a bypass of
the MAL dehydrogenase reaction via PYR, catalyzed by malic enzyme and
PYR carboxylase. Again, the result of this flux through the so-called
PYR shunt is net loss of one ATP, compared to the direct conversion of
MAL to OAA. The activity of the PYR shunt is mostly rather low, with the exception of the fast-growing P-limited culture. In this particular case, the reversed MAL dehydrogenase flux actually concludes a futile
cycle (Fig. 5).
An important aspect of flux estimate interpretation is the extent to
which the available data actually determine intracellular fluxes. To
ensure that indeed a global error minimum was identified, the parameter
search was started from at least five different starting points for
each flux calculation. These solutions were always similar, and the
flux solutions with the lowest 2 values are
presented in Fig. 4 and 5. These 2 values were
119, 153, and 222 at low D and 151, 117, and 210 at high
D for C, N, and P limitation experiments, respectively. Typically the 95% level of the 2 significance
test in such type of experiments is around 120 (9). Hence,
the identified 2 values are remarkably good
for the analysis of a biological system (65) and indicate
therefore a reliable description of the metabolic processes in B. subtilis.
The presented flux solutions were then subjected to further statistical
error analysis based on a linear approximation of the model around the
optimum flux solution (9, 65). For most fluxes we obtained
68% confidence intervals that were less than 10% of the estimated
flux (data not shown). One exception were fluxes in the PEP-PYR-OAA
triangle. The employed linearized statistical model is, however, not
well suited to assess the statistical relevance of these particular
fluxes, since the thus calculated confidence regions are much larger
than those obtained by other approaches (9). Based on the
almost-identical flux estimates that were obtained in five independent
calculations, it appears, therefore, that the confidence intervals for
the estimated fluxes in this triangle are similar to those of the other
fluxes in the network.
A second exception with a higher confidence interval was the oxidative
PP pathway flux for which 68% confidence intervals of about 25% were
calculated. In the present analysis, the calculated transhydrogenase
flux from NADPH to NADH (and vice versa) depends on the reported and
well-established cofactor dependencies (10, 16, 18) in the
biochemical reaction network (Fig. 4 and 5). As a consequence of higher
confidence intervals for the oxidative PP pathway flux, the
transhydrogenase flux is likewise not well determined. The net flux
from NADPH to NADH via transhydrogenase, however, is with 68%
confidence above 25% in all cases, except for the slow-growing,
P-limited culture where this net flux is at least 56%. When the
transhydrogenase reaction is omitted from the network, the
2 values of the corresponding flux solution
increase significantly from 119 to 167, 153 to 173, and 222 to 237 under C, N, and P limitations, respectively, at the low D.
This indicates that indeed transhydrogenase fluxes from NADPH to NADH
are consistent with all data.
Intracellular ATP and ADP concentrations.
To investigate the
direct impact of reduced energetic efficiency on the state of energy
metabolism in our cultures, we determined the intracellular
concentrations of two central intermediates of energy metabolism, ATP
and ADP. The concentrations of both adenine nucleotides increased
linearly with D in C-limited culture (Fig.
7). The ATP-to-ADP ratio that can be
calculated from these values was about 2 but should be treated with
care because the ADP concentrations have inherently large systematic
errors, since they were determined as the difference in fluorescence
between ATP and ATP plus ADP. The concentrations of ATP and ADP in
N-limited culture were very similar to those of C-limited cultures
(data not shown). As a consequence of the severe P limitation, the ATP and ADP concentrations in P-limited cultures were less than half of
those found in C- and N-limited cultures (data not shown). These data
show clearly that intracellular ATP concentrations depend on both the
specific growth rate and the environmental conditions in B. subtilis chemostat cultures. Although ATP-to-ADP ratios are not
well determined by the applied methodology, we have no indication that
these ratios differ significantly in the investigated cultures.
View larger version (25K):
[in this window]
[in a new window]
|
FIG. 7.
Intracellular concentration of ATP (circles) and ADP
(squares) in glucose-limited chemostat culture of B.
subtilis.
|
|
| |
DISCUSSION |
Glucose catabolism in B. subtilis.
The TCA
cycle in B. subtilis is subject to transcriptional control,
since CcpA- and CcpC-dependent carbon catabolite repression (54) reduces the expression of several TCA cycle genes in
the presence of glucose or other well-metabolized carbon sources
(18, 21, 58). In C-limited B. subtilis
chemostat culture, however, residual-glucose concentrations are
apparently not sufficient to trigger or fully activate this regulatory
response because TCA cycle fluxes are usually high (9, 46,
47). Despite residual-glucose concentrations of 0.8 and 2.4 g/liter, N-limited chemostat cultures exhibited high TCA cycle fluxes
that were similar to those seen in C-limited culture. Thus, the
apparent absence of catabolite repression in N-limited culture is
probably due to the absence of glutamate and/or glutamine from
the medium, which are synergistically required with glucose for full
repression of TCA cycle enzymes (17, 39).
Despite the absence of glutamate and glutamine, TCA cycle fluxes were
six- to eightfold lower in P-limited culture, with residual glucose
concentrations of 2 to 7 g/liter, compared to C- or N-limited culture
(Fig. 4 and 5). Although catabolite repression may play a role in this
deactivation of the TCA cycle, CcpA-mediated transcriptional activation
of genes involved in overflow metabolism (54) contributes probably significantly to these low TCA cycle fluxes by reducing the
availability of PYR and ACA for fluxes into the TCA cycle. This view is
supported by the extensive overflow metabolism in P-limited culture but
not in N- or C-limited culture (Fig. 3 to 5).
In addition to transcriptional regulation, several TCA cycle enzymes
are responsive to the energetic state of the cell (33), most prominently the major isoform of citrate synthase, which is a key
enzyme of the cycle and is competitively inhibited by ATP
(20). If ATP-dependent activation or reduced inhibition of
enzymes contributes significantly to the control of TCA cycle activity,
one would expect high TCA cycle fluxes in P-limited culture with
twofold-lower intracellular ATP concentrations than those in C- or
N-limited cultures. This is apparently not the case (Fig. 4 and 5), and
thus it appears that indeed translational control and not ATP-dependent
enzyme activity control is primarily responsible for down regulation of
TCA cycle activity under P limitation. It is currently unclear why TCA
cycle fluxes are high in N-limited culture and low in P-limited
culture, but one explanation may be that intracellular levels of
glutamate and glutamine are low under N limitation, so that
CcpA-dependent control is not fully activated under this condition.
Residual glucose concentration in N-limited cultures was probably
sufficient, however, to activate expression of overflow genes
(54), thus explaining intermediate acetate formation of
these cultures (Fig. 3B).
Based on a different methodology for metabolic flux analysis that
relied on significantly fewer NMR data, our lab estimated previously
high oxidative PP pathway fluxes in glucose-limited chemostat cultures
of B. subtilis (46). Although these experiments were done with a different strain, the results presented here (Fig. 4
and 5) and in a previous paper (9) suggest that oxidative PP pathway fluxes are usually less than 50% of the total glucose flux
in chemostat cultures of B. subtilis. When environmental conditions cause a very strong uncoupling of anabolism and catabolism, such as shown here for the slow-growing, P-limited culture, the oxidative PP pathway flux may, however, exceed 50% (Fig. 4).
Consequently, high transhydrogenase fluxes are needed to balance the
reducing equivalents NADPH and NADH. Although in vitro transhydrogenase activity was observed in B. subtilis (10),
conversion of NADPH into NADH may likewise be achieved by a carbon flux
cycling through isoenzymes with different cofactor specificity, i.e.,
the glyceraldehyde 3-phosphate dehydrogenases (14) or
malic enzyme (12). However, a significant contribution of
the latter reaction is unlikely because it would also affect the
13C-labeling pattern.
Cellular ATP balances.
The presented flux results provide a
holistic perspective on carbon metabolism and thus allow to construct
an ATP balance that quantifies partitioning of the total energy flux in
B. subtilis into different energy-consuming fluxes. To
account also for the incorporation of glucose into biomass and
metabolic (by-)products, all carbon fluxes were quantitatively
converted into ATP equivalents as outlined before. The energetic
equivalent of 1 mol of glucose in B. subtilis are 15 mol of
ATP equivalents, which corresponds to a maximum P-to-O (ATP per oxygen)
ratio of unity for respiratory ATP generation. Figure
8 depicts the partitioning of ATP
equivalents into the cellular processes in B. subtilis
considered here.
View larger version (32K):
[in this window]
[in a new window]
|
FIG. 8.
ATP balances of B. subtilis chemostat
cultures under carbon-limited (A and D), nitrogen-limited (B and E), or
phosphate-limited (C and F) conditions at D = 0.1 h1 (A to C) and D = 0.4 h1 (D to F). Balances are based on the total cellular
energy flux in each culture, which are the specific glucose uptake
rates of Fig. 3 multiplied by the maximum ATP that may be generated
from glucose at a P-to-O ratio of unity, i.e., 15. The partioning of
ATP equivalents into different metabolic processes includes the ATP
equivalents of carbon-containing end products. The following processes
were considered (in clockwise orientation): biomass formation (white);
riboflavin biosynthesis (dark gray); overflow metabolism (light gray),
i.e., the formation of acetate, diacyl, and acetoin; metabolic shifts
from optimal pathway usage (black), including fluxes in the futile
cycles of Fig. 6 and the PP pathway; maintenance metabolism (grid); and
excess ATP (hatched), which indicates the remaining ATP equivalents of
the total available in glucose after subtraction of the aforementioned
ATP-consuming processes.
|
|
Generally, biomass formation was the most costly process and required
between 40 and 70% of the available ATP equivalents, with the
exception of the slow-growing P-limited culture. These energy
requirements are strictly coupled to biosynthesis and polymerization and thus can be calculated in a straightforward fashion (10, 29,
45, 53). Recombinant riboflavin production accounted for only 2 to 4% of the total energy flux. The energetic relevance of overflow
metabolism was low under C limitation and moderate under N limitation
(Fig. 8). Under P limitation, however, overflow metabolism became the
primary dissipating flux of ATP equivalents in B. subtilis.
From the carbon fluxes presented in Fig.4 and 5 one can calculate
dissipation of ATP equivalents via futile cycling in PEP carboxykinase
and via metabolic shifts from energetically efficient to less efficient
pathways, i.e., fluxes through the oxidative PP pathway and the PYR
shunt. In contrast to what may be expected, dissipation of ATP
equivalents via these metabolic shifts or futile cycles was not
increased under excess-C conditions and the contribution to the ATP
balance was below 4% in all cultures investigated (Fig. 8). Although
the present flux analysis cannot quantify all potential futile cycles
(e.g., glucokinase/glucose-6-phosphatase or
phosphofructokinase/fructose-1,6-bisphosphatase), we found no evidence
for significantly increased activity of futile cycles under excess-C conditions.
Lastly we considered dissipation of ATP equivalents in maintenance
metabolism, which includes all energy expenditures that are not
directly related to growth (40). Based on the maintenance coefficient of 0.66 mmol of glucose per g of biomass per h that was
determined for glucose-limited, riboflavin-producing B. subtilis (47), one obtains a maintenance ATP
requirement of 9.9 mmol of ATP per g of biomass per h at the assumed
P/O of 1 for C-limited cultures. This value was also used for the
excess-C cultures. The maintenance metabolism thus calculated becomes a
major ATP-dissipating process at low D but contributes not
more than 10% to the ATP balance at high D (Fig. 8). The
assumed maintenance coefficient is significantly higher than the one
obtained recently for the strain used here (10). These
experiments were, however, not designed to accurately determine this
coefficient, and newer data suggest that maintenance metabolism in the
strain used here is indeed as high as was assumed above (N. Zamboni and
U. Sauer, unpublished data).
The sum of all considered ATP equivalent-consuming processes in the
slow-growing C-limited chemostat culture matched the available ATP
equivalents from glucose exactly (Fig. 8). This demonstrates that all
energetically relevant cellular processes were comprehensively considered in this particular case. Under all other conditions, however, a remarkably constant fraction of about 20% of the available ATP equivalents cannot be assigned to a specific ATP-consuming flux and
is therefore referred to as excess ATP (Fig. 8). This value is an upper
bound for the potential energetic contribution of all other
ATP-dissipating processes that cannot be assessed from the present
data, for example ion leakage (7, 40), nonquantified futile enzyme cycles, or growth rate-dependent maintenance requirements (40). The similarity of the estimated excess ATP fraction
in C-limited and excess-C cultures at high D (Fig. 8D to F),
suggests that there are no additional ATP-dissipating mechanisms that
operate specifically under excess-C conditions in fast-growing B. subtilis. In excess-C cultures at low D, however, about
20% of the total energy flux may be dissipated by mechanisms that are
specific to C sufficiency.
Any conclusion on cellular energetics depends to some extent on the
assumed P/O. The above conclusions, however, are not critically dependent on a P/O of unity, since changes in P/O affect ATP production and consumption similarly. Specifically, the excess ATP equivalent values (Fig. 8) would be maximally 3% higher or lower when P/O varied
within the biologically reasonable range for B. subtilis of
0.8 to 1.33, respectively. This P/O range is biologically reasonable because metabolic NADH is primarily oxidized via the
aa3 oxidase branch (translocation of
four protons per NADH) under the present conditions (25, 27, 42,
66, 67) and the H+-to-ATP ratio of ATP
synthase is firmly established to be 4 (13, 43). Already a
P/O of 1.33 would be unlikely, since it would, for example, require
that the H+-to-ATP ratio of ATP synthase was only
3. In most cases, excess ATP would disappear at P/O between 0.3 and
0.4. Such P-to-O ratios, however, appear to be unrealistically low, so
that probably the majority of the excess ATP fraction is indeed
dissipated by processes that are not being considered at present.
The presented new concept for a cellular ATP balance that
comprehensively considers the total energy flux from both carbon and
ATP metabolism enables quantitative insights into cellular energy
management. In particular, this concept attributes the previously
observed discrepancy between theoretical and experimentally observed
maximum biomass yields on ATP (see, for example, references 31,
53, and 62) to specific energy-dissipating processes.
Financial support was obtained through a scholarship from the
Boehringer Ingelheim Fonds to M.D.
| 1.
|
Bäck, T., and H.-P. Schwefel.
1993.
An overview of evolutionary algorithms for parameter optimization.
Evol. Comp.
1:1-23.
|
| 2.
|
Bakker, E. P.
1993.
Alkali cation transport systems in prokaryotes.
CRC Press, Boca Raton, Fla.
|
| 3.
|
Bergmeyer, H. U.
1985.
Methods of enzymatic analysis, vol. IV.
VCH Publisher, Deerfield Beach, Fla.
|
| 4.
|
Brooke, A. C.,
M. M. Attwood, and D. W. Tempest.
1990.
Metabolic fluxes during the growth of thermotolerant methylotrophic Bacillus strains in methanol-sufficient chemostat cultures.
Arch. Microbiol.
153:591-595[CrossRef].
|
| 5.
|
Chambost, J. P., and D. G. Fraenkel.
1980.
The use of 6-labeled glucose to assess futile cycling in Escherichia coli.
J. Biol. Chem.
255:2867-2869[Abstract/Free Full Text].
|
| 6.
|
Chao, Y.-P., and J. C. Liao.
1994.
Metabolic responses to substrate futile cycling in Escherichia coli.
J. Biol. Chem.
269:5122-5126[Abstract/Free Full Text].
|
| 7.
|
Cook, G. M., and J. B. Russell.
1994.
Energy-spilling reactions of Streptococcus bovis and resistance of its membrane to proton conductance.
Appl. Environ. Microbiol.
60:1942-1948[Abstract/Free Full Text].
|
| 8.
|
Daldal, F., and D. G. Fraenkel.
1983.
Assessment of a futile cycle involving reconversion of fructose 6-phosphate to fructose 1,6-bisphosphate during gluconeogenic growth of Escherichia coli.
J. Bacteriol.
153:390-394[Abstract/Free Full Text].
|
| 9.
|
Dauner, M.,
J. E. Bailey, and U. Sauer.
2001.
Metabolic flux analysis with a comprehensive isotopomer model in Bacillus subtilis.
Biotechnol. Bioeng.
76:144-156[CrossRef][Medline].
|
| 10.
|
Dauner, M., and U. Sauer.
2001.
Stoichiometric growth model for riboflavin-producing Bacillus subtilis.
Biotechnol. Bioeng.
76:132-143[CrossRef][Medline].
|
| 11.
|
Dawson, P. S. S.
1985.
Continuous cultivation of microorganisms.
Crit. Rev. Biotechnol.
2:315-372.
|
| 12.
|
Diesterhaft, M. D., and E. Freese.
1973.
Role of pyruvate carboxylase, phosphoenolpyruvate carboxykinase, and malic enzyme during growth and sporulation of Bacillus subtilis.
J. Biol. Chem.
248:6062-6070[Abstract/Free Full Text].
|
| 13.
|
Fillingame, R. H.
1997.
Coupling H+ transport and ATP synthesis in F1 F0-ATP synthases: glimpses of interacting parts in a dynamic molecular machine.
J. Exp. Biol.
200:217-224[Abstract].
|
| 14.
|
Fillinger, S.,
S. Boschi-Muller,
S. Azza,
E. Dervyn,
G. Branlant, and S. Aymerich.
2000.
Two glyceraldehyde 3-phosphate dehydrogenases with opposite physiological roles in a non-photosynthetic bacterium.
J. Biol. Chem.
275:14031-14037[Abstract/Free Full Text].
|
| 15.
|
Gerhardt, P.,
R. G. E. Murray,
W. A. Wood, and N. R. Krieg.
1994.
Methods for general and molecular bacteriology.
ASM Press, Washington, D.C.
|
| 16.
|
Gottschalk, G.
1986.
Bacterial metabolism, 2nd ed.
Springer-Verlag, New York, N.Y.
|
| 17.
|
Hanson, R. S., and D. P. Cox.
1967.
Effect of different nutritional conditions on the synthesis of tricarboxylic acid cycle enzymes.
J. Bacteriol.
93:1777-1787[Abstract/Free Full Text].
|
| 18.
|
Hederstedt, L.
1993.
The Krebs citric acid cycle, p. 181-197.
In
A. L. Sonenshein, J. A. Hoch, and R. Losick (ed.), Bacillus subtilis and other gram-positive bacteria: biochemistry, physiology, and molecular genetics. American Society for Microbiology, Washington, D.C.
|
| 19.
|
Hollywood, N., and H. W. Doelle.
1976.
Effect of specific growth rate and glucose concentration on growth and glucose metabolism of Escherichia coli K-12.
Microbios
17:23-33[Medline].
|
| 20.
|
Jin, S., and A. L. Sonenshein.
1996.
Characterization of the major citrate synthase of Bacillus subtilis.
J. Bacteriol.
178:3658-3660[Abstract/Free Full Text].
|
| 21.
|
Jourlin-Castelli, C.,
N. Mani,
M. M. Nakano, and A. L. Sonenshein.
2000.
CcpC, a novel regulator of the LysR family required for glucose repression of the citB gene in Bacillus subtilis.
J. Mol. Biol.
295:865-878[CrossRef][Medline].
|
| 22.
|
Katz, J., and R. Rognstad.
1978.
Futile cycling in glucose metabolism.
Trends Biochem. Sci.
3:171-174.
|
| 23.
|
Lang, W. K.,
K. Glassey, and A. R. Archibald.
1982.
Influence of the phosphate supply on teichoic and teichuronic acid content of Bacillus subtilis cell walls.
J. Bacteriol.
151:367-375[Abstract/Free Full Text].
|
| 24.
|
Larsson, C.,
U. von Stockar,
I. Marrison, and L. Gustafsson.
1993.
Growth and metabolism of Saccharomyces cerevisiae in chemostat cultures under carbon-, nitrogen-, or carbon- and nitrogen-limiting conditions.
J. Bacteriol.
175:4809-4816[Abstract/Free Full Text].
|
| 25.
|
Lauraeus, M., and M. Wikström.
1993.
The terminal quinol oxidases of Bacillus subtilis have different energy conservation properties.
J. Biol. Chem.
268:11470-11473[Abstract/Free Full Text].
|
| 26.
|
Lifely, M. R.,
E. Tarelli, and J. Baddiley.
1980.
The teichuronic acid from the wall of Bacillus licheniformis ATCC 9945.
Biochem. J.
191:305-318[Medline].
|
| 27.
|
Liu, X., and H. W. Taber.
1998.
Catabolite regulation of the Bacillus subtilis ctaBCDEF gene cluster.
J. Bacteriol.
180:6154-6173[Abstract/Free Full Text].
|
| 28.
|
Martins, M. L. L., and D. W. Tempest.
1991.
Metabolic response of Bacillus stearothermophilus chemostat cultures to a secondary oxygen limitation.
J. Gen. Microbiol.
137:1391-1396.
|
| 29.
|
Neidhardt, F. C.,
J. L. Ingraham, and M. Schaechter.
1990.
Physiology of the bacterial cell: a molecular approach.
Sinauer Associates, Inc., Sunderland, Mass.
|
| 30.
|
Neijssel, O. M.,
E. T. Buurman, and M. J. Teixeira de Mattos.
1990.
The role of futile cycles in the energetics of bacterial growth.
Biochim. Biophys. Acta
1018:252-255[Medline].
|
| 31.
|
Neijssel, O. M.,
M. J. Teixeira de Mattos, and D. W. Tempest.
1996.
Growth yield and energy distribution, p. 1683-1692.
In
F. C. Neidhardt, R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella: cellular and molecular biology, 2nd ed. ASM Press, Washington, D.C.
|
| 32.
|
Neijssel, O. M., and D. W. Tempest.
1979.
The physiology of metabolite over-production.
Symp. Soc. Gen. Microbiol.
29:53-82.
|
| 33.
|
Ohné, M.
1975.
Regulation of the dicarboxylic acid part of the citric acid cycle in Bacillus subtilis.
J. Bacteriol.
122:224-234[Abstract/Free Full Text].
|
| 34.
|
Pavlik, J. G., and H. J. Rogers.
1973.
Selective extraction of polymers from cell walls of Gram-positive bacteria.
Biochem. J.
131:619-621[Medline].
|
| 35.
|
Pennock, J., and D. W. Tempest.
1988.
Metabolic and energetic aspects of the growth of Bacillus stearothermophilus in glucose-limited and glucose-sufficient chemostat culture.
Arch. Microbiol.
150:452-459[CrossRef].
|
| 36.
|
Perkins, J. B.,
A. Sloma,
T. Hermann,
K. Theriault,
E. Zachgo,
T. Erdenberger,
N. Hannett,
N. P. Chatterjee,
V. Williams II,
G. A. Rufo, Jr.,
R. Hatch, and J. Pero.
1999.
Genetic engineering of Bacillus subtilis for the commercial production of riboflavin.
J. Ind. Microbiol. Biotechnol.
22:8-18.
|
| 37.
|
Petersen, S.,
A. A. de Graaf,
L. Eggeling,
M. Möllney,
W. Wiechert, and H. Sahm.
2000.
In vivo quantification of parallel and bidirectional fluxes in the anaplerosis of Corynebacterium glutamicum.
J. Biol. Chem.
275:35932-35941[Abstract/Free Full Text].
|
| 38.
|
Press, W. H.,
S. A. Teukolsky,
W. T. Vetterling, and B. P. Flannery.
1995.
Numerical recipes in C: the art of scientific computing.
Cambridge University Press, New York, N.Y.
|
| 39.
|
Rosenkrantz, M. S.,
D. W. Dingman, and A. L. Sonenshein.
1985.
Bacillus subtilis citB gene is regulated synergistically by glucose and glutamine.
J. Bacteriol.
164:155-164[Abstract/Free Full Text].
|
| 40.
|
Russell, J. B., and G. M. Cook.
1995.
Energetics of bacterial growth: balance of anabolic and catabolic reactions.
Microbiol. Rev.
59:48-62[Abstract/Free Full Text].
|
| 41.
|
Saier, M. H. J.,
M. J. Fagan,
C. Hoischen, and J. Reizer.
1993.
Transport mechanisms, p. 133-156.
In
A. L. Sonenshein, J. A. Hoch, and R. Losick (ed.), Bacillus subtilis and other gram-positive bacteria: biochemistry, physiology, and molecular genetics. American Society for Microbiology, Washington, D.C.
|
| 42.
|
Santana, M.,
F. Kunst,
M. F. Hullo,
G. Rapoport,
A. Danchin, and P. Glaser.
1992.
Molecular cloning, sequencing, and physiological characterization of the qox operon from Bacillus subtilis encoding the aa3-600 quinol oxidase.
J. Biol. Chem.
267:10225-10231[Abstract/Free Full Text].
|
| 43.
|
Saraste, M.
1999.
Oxidative phosphorylation at the fin de siecle.
Science
283:1488-1493[Abstract/Free Full Text].
|
| 44.
|
Sauer, U., and J. E. Bailey.
1999.
Estimation of P-to-O ratio in Bacillus subtilis and its influence on maximum riboflavin yield.
Biotechnol. Bioeng.
64:750-754[CrossRef][Medline].
|
| 45.
|
Sauer, U.,
D. C. Cameron, and J. E. Bailey.
1998.
Metabolic capacity of Bacillus subtilis for the production of purine nucleotides, riboflavin, and folic acid.
Biotechnol. Bioeng.
59:227-238[CrossRef][Medline].
|
| 46.
|
Sauer, U.,
V. Hatzimanikatis,
J. E. Bailey,
M. Hochuli,
T. Szyperski, and K. Wüthrich.
1997.
Metabolic fluxes in riboflavin-producing Bacillus subtilis.
Nat. Biotechnol.
15:448-452[CrossRef][Medline].
|
| 47.
|
Sauer, U.,
V. Hatzimanikatis,
H.-P. Hohmann,
M. Manneberg,
A. P. G. M. van Loon, and J. E. Bailey.
1996.
Physiology and metabolic fluxes of wild-type and riboflavin-producing Bacillus subtilis.
Appl. Environ. Microbiol.
62:3687-3696[Abstract].
|
| 48.
|
Sauer, U.,
D. R. Lasko,
J. Fiaux,
M. Hochuli,
R. Glaser,
T. Szyperski,
K. Wüthrich, and J. E. Bailey.
1999.
Metabolic flux ratio analysis of genetic and environmental modulations of Escherichia coli central carbon metabolism.
J. Bacteriol.
181:6679-6688[Abstract/Free Full Text].
|
| 49.
|
Schirawski, J., and G. Unden.
1998.
Menaquinone-dependent succinate dehydrogenase of bacteria catalyzes reversed electron transport driven by the proton potential.
Eur. J. Biochem.
257:210-215[Medline].
|
| 50.
|
Schmidt, K.,
L. C. Nørregaard,
B. Pedersen,
A. Meissner,
J. Ø. Duus,
J. O. Nielsen, and J. Villadsen.
1999.
Quantification of intracellular metabolic fluxes from fractional enrichment and 13C-13C coupling constraints on the isotopomer distribution in labeled biomass components.
Metab. Eng.
1:166-179[CrossRef][Medline].
|
| 51.
|
Schreier, H. J.
1993.
Biosynthesis of glutamine and glutamate and the assimilation of ammonia, p. 281-298.
In
A. L. Sonenshein, J. A. Hoch, and R. Losick (ed.), Bacillus subtilis and other gram-positive bacteria: biochemistry, physiology, and molecular genetics. American Society for Microbiology, Washington, D.C.
|
| 52.
|
Snay, J.,
J. W. Jeong, and M. M. Ataai.
1989.
Effects of growth conditions on carbon utilization and organic by-product formation in B. subtilis.
Biotechnol. Progr.
5:63-69.
|
| 53.
|
Stouthamer, A. H.
1979.
The search for correlation between theoretical and experimental growth yields, p. 1-48.
In
J. R. Quayle (ed.), Microbial biochemistry, vol. 21. University Park Press, Baltimore, Md.
|
| 54.
|
Stülke, J., and W. Hillen.
2000.
Regulation of carbon catabolism in Bacillus species.
Annu. Rev. Microbiol.
54:849-880[CrossRef][Medline].
|
| 55.
|
Szyperski, T.
1995.
Biosynthetically directed fractional 13C-labeling of proteinogenic amino acids: an efficient analytical tool to investigate intermediary metabolism.
Eur. J. Biochem.
232:433-448[Medline].
|
| 56.
|
Szyperski, T.,
R. W. Glaser,
M. Hochuli,
J. Fiaux,
U. Sauer,
J. E. Bailey, and K. Wüthrich.
1999.
Bioreaction network topology and metabolic flux ratio analysis by biosynthetic fractional 13C-labeling and two-dimensional NMR spectroscopy.
Metab. Eng.
1:189-197[CrossRef][Medline].
|
| 57.
|
Teixeira de Mattos, M. J., and O. M. Neijssel.
1997.
Bioenergetic consequences of microbial adaptation to low-nutrient environments.
J. Biotechnol.
59:117-126[CrossRef][Medline].
|
| 58.
|
Tobisch, S.,
D. Zühlke,
J. Bernhardt,
J. Stülke, and M. Hecker.
1999.
Role of CcpA in regulation of the central pathways of carbon catabolism in Bacillus subtilis.
J. Bacteriol.
181:6996-7004[Abstract/Free Full Text].
|
| 59.
|
Torres, J. C.,
V. Guixé, and J. Babul.
1997.
A mutant phosphofructokinase produces a futile cycle during gluconeogenesis in Escherichia coli.
Biochem. J.
327:675-684.
|
| 60.
|
Tran, Q. H., and G. Unden.
1998.
Changes in the proton potential and the cellular energetics of Escherichia coli during growth by aerobic and anaerobic respiration or by fermentation.
Eur. J. Biochem.
251:538-543[Medline].
|
| 61.
|
Trumpower, B. L., and R. B. Gennis.
1994.
Energy transduction by cytochrome complexes in mitochondrial and bacterial respiration: The enzymology of coupling electron transfer reactions to transmembrane proton translocation.
Annu. Rev. Biochem.
63:675-716[Medline].
|
| 62.
|
Walsh, K., and D. E. Koshland, Jr.
1984.
Determination of flux through the branch point of two metabolic cycles.
J. Biol. Chem.
259:9646-9654[Abstract/Free Full Text].
|
| 63.
|
Wiechert, W.
2001.
13C metabolic flux analysis.
Metab. Eng.
3:195-206[CrossRef][Medline].
|
| 64.
|
Wiechert, W., and A. A. de Graaf.
1997.
Bidirectional reaction steps in metabolic networks. I. Modeling and simulation of carbon isotopes labeling experiments.
Biotechnol. Bioeng.
55:101-117[CrossRef].
|
| 65.
|
Wiechert, W.,
C. Siefke,
A. A. de Graaf, and A. Marx.
1997.
Bidirectional reaction steps in metabolic networks. II. Flux estimation and statistical analysis.
Biotechnol. Bioeng.
55:118-135[CrossRef].
|
| 66.
|
Winstedt, L., and C. von Wachenfeldt.
2000.
Terminal oxidases of Bacillus subtilis strain 168: one quinol oxidase, cytochrome aa3 or cytochrome bd, is required for aerobic growth.
J. Bacteriol.
182:6557-6564[Abstract/Free Full Text].
|
| 67.
|
Winstedt, L.,
K.-I. Yoshida,
Y. Fujita, and C. von Wachenfeldt.
1998.
Cytochrome bd biosynthesis in Bacillus subtilis: characterization of the cydABCD operon.
J. Bacteriol.
180:6571-6580[Abstract/Free Full Text].
|
| 68.
|
Yee, L., and H. W. Blanch.
1993.
Defined media optimization for growth of recombinant Escherichia coli X90.
Biotechnol. Bioeng.
41:221-230[CrossRef].
|
![[Feedback]](/icons/banner/feedbackACT.gif){kind=icon}
![[For Subscribers]](/icons/banner/subscriberACT.gif){kind=icon}
![[Archive]](/icons/banner/archiveACT.gif){kind=icon}
![[Search]](/icons/banner/searchACT.gif){kind=icon}
![[Contents]](/icons/banner/tocACT.gif){kind=icon}
Top
Abstract
Introduction
