In the earlier phase, both glc and glu were utilized, resulting in average ATP and ADP concentrations in cells of 0.50 ± 0.17 lmolÆg1of dry cell weight DCW and an undetermined level ADP
Trang 1P NMR studies of energy metabolism in
Yasushi Noguchi1, Nobuhisa Shimba2, Yoshio Kawahara1, Ei-ichiro Suzuki2and Shinichi Sugimoto1
1
Fermentation & Biotechnology Laboratories and2Central Research Laboratories, Ajinomoto Co., Inc., Kawasaki-ku,
Kawasaki, Kanagawa, Japan
Corynebacterium ammoniagenes is an overproducer of
xanthosine-5¢-monophosphate (XMP) by consuming either
glcose (glc) or glutamic acid (glu) Its energy metabolism was
studied in vivo using31P NMR spectroscopy coupled with a
circulating fermentation system (CFS) CFS enabled us to
validate directly the cellular dependency on carbon sources
and changes in biomolecules produced according to
altera-tions in the cellular energetic status For the most efficient
XMP production, the glutamic acid and glcose molar ratios
(glu/glc) in the medium were adjusted to a molar ratio of
0.31 The31P NMR illustrated the two distinct phases of the
cellular energetic status due to the availability of the
sub-strates from the medium In the earlier phase, both glc
and glu were utilized, resulting in average ATP and ADP
concentrations in cells of 0.50 ± 0.17 lmolÆg)1of dry cell
weight (DCW) and an undetermined level
ADP concentration in the later phase increased to 2.15 ± 1.30 lmolÆg)1of DCW, while the ATP concentra-tion decreased to an undetectable level in associaconcentra-tion with a remarkable decrease in XMP production This decrease in the XMP-producing ability was associated with an increase
in production of the by-product hypoxanthine Because glu was found to be consumed completely during the earlier phase, glc was the only available substrate in the later phases
These findings by in vivo NMR indicate that changes in the carbon metabolism profoundly affect XMP production by
C ammoniagenes
Keywords: xanthosine-5¢-monophosphate; in vivo NMR; energy metabolism; Corynebacterium ammoniagenes
In order to understand the microbial production of purine
and other nucleotides, evaluation of the cellular energy
metabolism is extremely important because the nucleotide
biosynthesis requires high levels of ATP [1,2] Difficulties in
obtaining the energy metabolism profile from living cells
are details of the regulatory process of nucleotide
biosyn-thesis which remain to be studied NMR spectroscopy has
allowed in vivo measurement of metabolite concentrations,
thereby permitting assessment of the dynamic changes in
metabolic pathways and cellular regulatory mechanisms
[3,4] The circulating fermentation system (CFS) that we
have previously developed [5] enables us to prolong an
NMR spectroscopic measurement period under various
culture conditions
Corynebacterium ammoniagenes is a Gram-positive,
coryneform bacterium important to the industrial
pro-duction of flavor-enhancing purine nucleotides such as
inosine-5¢-monophosphate and
xanthosine-5¢-monophos-phate [6–8] Several studies have reported adenine
and guanine auxotrophic mutants of C ammoniagenes ATCC6872 that excessively secrete purine nucleotides into their cultures [6–8] In these studies, the balance between the oxidative pentose phosphate (PP) cycle for supplementation
of the carbon skeleton and the tricarboxylic acid (TCA) cycle for maximization of ATP production was discussed However, direct estimation of contribution of these two metabolic pathways in the nucleotide production was not available
In the present study, we investigate the cellular energy metabolism in XMP-overproducing C ammoniagenes, monitor phosphate-containing metabolites by using 31P NMR spectroscopy, and discuss cellular energetics in nucleotide production
Experimental procedures
Chemicals MDP (methylene diphosphonic acid) was purchased from Sigma Chemical Co (St Louis, MO, USA) All other chemicals were commercially available and of the highest grade
Bacterial strain and cultivation conditions XMP-overproducing C ammoniagenes is an adenine and guanine auxotrophic mutant isolated from the wild-type strain C ammoniagenes ATCC6872 For in vivo NMR studies, the inoculum was prepared on 50 plates of the modified LB medium, which was composed of 10 gÆL)1
Correspondence to Y Noguchi, Fermentation & Biotechnology
Laboratories, Ajinomoto Co., Inc., Suzuki-cho 1-1, Kawasaki-ku,
Kawasaki, Kanagawa 210-8681, Japan.
Fax: + 81 44 2117609, Tel.: + 81 44 2105898,
E-mail: yasushi_noguchi@ajinomoto.com
Abbreviations: CFS, circulating fermentation system; DCW, dry cell
weight; MDP, methylene diphosphonic acid; PP, pentose phosphate;
TCA, tricarboxylic acid cycle; XMP, xanthosine-5¢-monophosphate.
(Received 15 November 2002, revised 23 February 2003,
accepted 25 April 2003)
Trang 2tryptone, 5 gÆL)1yeast extract, 5 gÆL)1NaCl, and 20 gÆL)1
bacto-agar plus 0.01 gÆL)1adenine and 0.01 gÆL)1guanine
For in vivo NMR experiments, fermentations were carried
out in a fermentor with an initial culture volume of 700 mL
at 30C and pH 6.5 XMP-overproducing strains had
been allowed to grow in the above medium supplemented
with 100 gÆL)1 glc, 5 gÆL)1 K2HPO4, 5 gÆL)1 KH2PO4,
0.05 gÆL)1 adenine, 0.05 gÆL)1 guanine, 1 gÆL)1 yeast
extract, 0.5 gÆL)1 NaCl, 0.25 gÆL)1 CaCl2, 1 gÆL)1
MgSO4Æ4H2O, and 0.05 mg L)1FeSO4Æ7H2O To optimize
medium glu/glc ratio to gain maximum XMP production,
0, 12.5, 25, 37.5 and 50 gÆL)1glu, respectively, were added
to the above medium The C ammoniagenes wild type and
Escherichia coli W3110 were cultured under the same
culture conditions
Fermentation system
For the in vivo NMR measurements, a previously
construc-ted CFS was used [5] In our system, the agitation speed of
the fermentor was automatically regulated to maintain the
dissolved oxygen tension (DOT) values in the fermentation
vessel Oxygen and carbon dioxide consumption rates were
measured using an exhaust gas analyzer system (Able Co.,
Tokyo, Japan) The temperature of the whole system was
kept at 30C using a circulating water bath, and
acidifica-tion was corrected by automated addiacidifica-tions of 10%
NH4OH
NMR operation
Using cells at high density (> 5 g of DCWÆL)1),31P NMR
signals were recorded at 161 MHz with a Bruker DSX
400 WB spectrometer, where 4-k data points were recorded
with 1280 transients and a spectral width of 16 kHz The
spectra were typically acquired with the following
param-eters: pulse width, 34 ls (90 flip angle); repetition time,
1.5 s To enhance the resolution, the free induction decay
was multiplied by an exponential window function prior to
Fourier transformation To quantify the intracellular
metabolites, MDP was used as a concentration standard
in the NMR Partially relaxed MDP was used to estimate
metabolite concentrations following reported methods [5,9]
Analysis
The DCW was determined
(D) at 660 nm using a Shimadzu UV260 spectroscope
and comparison with an optical density vs dry weight
calibration curve (the coefficient¼ 2.5) The concentrations
of glc and glu were determined enzymatically using a biotech-analyzer (Asahi Kasei Co., Tokyo, Japan) The concentrations of XMP and hypoxanthine in the culture were assayed by the HPLC (high-performance liquid chromatography) method as described previously [10,11]
Results and discussion
XMP production byC ammoniagenes
To validate the performance of C ammoniagenes, an XMP-overproducing strain, cells were batch-cultured as described
in the Experimental procedures, where both glc and glu were used as carbon sources Both concentrations in the growing medium were optimized to gain maximum XMP production (Table 1) As shown in Table 1, an increase in the glu/glc ratio (glu/glc) induced a significant increase in XMP production, and this increase in XMP production was coupled with a reduction in production of the by-product, hypoxanthine XMP production attained nearly plateau levels at a glu/glc molar ratio of 0.31 Further increases in glu/glc ratio to 0.61 lowered the XMP level (Table 1) Thus, a 0.31 molar ratio of the glu/glc was standardized for the in vivo 31P NMR measurements to validate the effect of glu on XMP production Changes in D
with glc, glu, XMP, and hypoxanthine concentrations in this culture condition are shown in Fig 1 Under this experimental condition, 68.9 mM XMP and 28.9 mM
hypoxanthine were obtained at the end of cultivation (Fig 1B) A rapid increase in the specific XMP produc-tion rate in parallel with the cell growth was observed until 25 h of the culturing period, but the specific XMP production rate then drastically decreased when glu in the growing medium was thoroughly consumed (Fig 1C) Contrary to XMP, hypoxanthine was not accumulated during the glu-consuming phase (phase I), but much accumulated during the glu-deficient phase (phase II), suggesting that hypoxanthine production was induced through a switch in the carbon flux In Table 1, glu increased the XMP/Hyp ratio, being dependent on the increased level of glu/glc ratio up to 0.46, which mainly resulted from the extension of the length of phase I Additionally, a noticeable reduction in either O2 con-sumption or CO2 production rates occurred during phase II, which coincided with the onset of hypoxanthine
5
production (Fig 1D) Cumulatively, it can be concluded that glu availability determines the efficiency of energy production or TCA cycle fluxes, which may specify Table 1 Effect of glutamic acid on XMP and hypoxanthine (Hyp)production in C ammoniagenes XMP overproducer.
concentrations were assayed as described in Experimental procedures The dry cell weight (DCW) was determined by comparison with a D vs dry weight calibration curve.
7
Glu
(mol)
Glc (mol)
Glu/glc (mol/mol)
XMP (mM)
Hyp
DCW (gÆL)1)
Trang 3whether primarily XMP or hypoxanthine production
takes place
Energetic status during the XMP production phase
To understand further the regulatory mechanism of
nucleotide production, dependency of the cellular
ener-getic on the carbon source was investigated by measuring
cellular ATP and ADP by the CFS system coupled with
31P NMR In Fig 2, typical NMR spectra during the
phase of XMP production (phase I) are illustrated with
peak assignments based on the previously published data
[12,13] Under our experimental condition, sugar phos-phate and intracellular inorganic phosphos-phate (Pin
i ) signals
at approximately)5–0 p.p.m were not distinct from the accumulated XMP and inorganic phosphate added as
K2HPO4 and KH2PO4 for essential substrates for XMP production
During phase I, ATPb and ATPc plus ADPb signals were maintained at very low intensity levels During phase II (hypoxanthine producing phase), ATPb signals were unde-tectable, but the ATPc plus ADPb intensity increased fivefold in contrast to those during phase I Chronological changes XMP producers, i.e cellular ATP, ADP and
Fig 1 Changes in growth, substrates, and products of C ammoniagenes in the XMP-production phase (A) Growth (j), residual glc (s), and residual glu (h) (B) XMP (j) and hypoxanthine (m) production (C) Glc (s) and glu (h) consumption, and XMP production rates (j) (D) Oxygen consumption (d) and CO 2 production (s) rates In phase I (P-I), the bacterium consumed glu in parallel with glc, and in phase II (P-II), the bacterium consumed glc as a sole substrate Cultivation was performed as described in the Experimental procedures.
Fig 2 Representative31P NMR spectra of metabolites from the XMP overproducer in phase I (A)and II (B) In spectra A and B, cell density during NMR observations were approximately 8 gÆL)1and 23 gÆL)1, respect-ively Abbreviations for resonances are: methylene diphosphonic acid (MDP), uridine diphosphate glc (UDP-glc), b and c phosphate
of adenine nucleotide phosphates (ATPb, and ADPb plus ATPc, respectively) The spectrum consisted of 1280 scans and was acquired at
161 MHz with a spectral width of 16 kHz, a 90 pulse angle, and a recycling time of 1.5 s Chemical shifts are given in p.p.m from 85% H PO
Trang 4NADP concentrations during the production phase are
shown in Fig 3 In phase I, ATP concentration was low but
above the detectable level, but ADP concentration was
undetectable ADP concentration increased in phase II,
while the ATP concentration, in turn, became undetectable
Cellular NADP as shown in Fig 3B was kept at similar
levels throughout the cultivation period Thus, substrate
availability seemed not to change NADP levels
By introducing C ammoniagenes wild-type strain
(ATCC6872) and E coli W3110, intracellular ATP and
ADP concentrations were measured to confirm whether the
low energetic status observed in this XMP producer could
be reproduced in these two wild-type strains under the
same culture condition Comparisons are made in Table 2
Between the two wild-type strains, cellular ATP plus ADP
levels and ATP/ADP ratios were not significantly different
each other through phases I and II Contrary, in the XMP
producer during phase I, the average ATP plus ADP
concentration, as well as ATP/ADP ratios, were much lower than those in the two wild-type strains (Table 2) ATP plus ADP concentration in the XMP producer, the mutant
C ammoniagenescorresponded to 14% of that in the wild type (Table 2) During phase II, the average ATP plus ADP concentration raised 58% of that in the wild-type strain, but ATP concentration further decreased to an undetectable level (Table 2) Thus, low ATP levels or ATP plus ADP concentrations continuously observed in the XMP-produ-cing phase is specific to this strain and this low energetic status will not be attributable to the unavailability of substrates, but to the specific metabolic characteristic of XMP production itself
A prominent enlargement in the ATP and ADP pool size from phase Ito IIin the XMP producer was associated with
a shift from ATP to ADP production This shift may be explained by an effect of glu deficiency on the central carbon flux, which specifically occurs during XMP production In fact, an increase in ATP concentration and ATP/ADP ratio could be induced during phase Iby increasing glu concen-tration in the culture medium (Fig 4) Dauner et al [14] have simulated a maximization of the flux in riboflavin production in Bacillus subtilis, and proposed the importance
of energy supplementation on which activities of the TCA cycle and the respiratory chain depend Several reports also
Fig 3 Changes in cellular ATP, ADP and ATP/ADP of the XMP
overproducer in the production phase (A) Cellular ATP (s) and ADP
(d) concentrations are shown (B) Cellular NADP (j) is shown To
quantify the intracellular metabolites, MDP was used as a
concen-tration standard in the NMR All data were normalized by DCW as
described in Experimental procedures.
Table 2 Comparison of cellular metabolites between the XMP-overproducing and wild-type strains Cultivation conditions are summarized in the Experimental procedures To quantify the intracellular metabolites, NADP was used as a concentration standard in the NMR Partially relaxed MDP
8,9 was used to estimate metabolite concentrations ND, not determined Values are the mean ± SD.
8,9
Strain
Average concentrations (pmolÆg)1of DCW)
C ammoniagenes
XMP overproducer
10
Fig 4 Effect of glu on intracellular ATP and ADP concentrations in the phase I Intracellular ATP and ADP concentrations in phase I were estimated based on 31 P NMR data The data shown are the mean ± SD.
Trang 5proposed the importance of the balance between TCA and
the oxidative PP cycles in nucleotide production [14–16]
These reports suggested that an enhancement of the net flux
of the oxidative PP cycle that supplied carbon skeletons led
to a reduction of ATP production due to a reduced TCA
cycle flux in nucleotide production [14–16] In our
experi-ment, XMP production decreased sharply during phase II,
simultaneous with an increased production of
hypoxan-thine, an XMP by-product (Fig 1) In association with this
reduction of XMP production, glc utilization was also
reduced, indicating that the reduced XMP flux through the
PP cycle affects glc oxidation This will result in reduction of
NADH generation rate from TCA cycle and a decrease in
ATP pool The reduced ATP availability seems to enhance
the production of the by-product, hypoxanthine This
cause–result corresponds well with in vivo NMR results in
this study, as well as previous reports [15–17] A
consider-able increase in ADP synthesis was observed during phase
II; the true reason for this increase remains to be elucidated
and whether biosynthesis of adenine nucleotides is
speci-fically enhanced or oxidative phosphorylation is specispeci-fically
reduced
This study represents the first trial of in vivo NMR
observation of bacterial nucleotide production Our results
demonstrate that the control of energy metabolism is crucial
for bacterial nucleotide production as, for instance,
main-tenance of efficient ATP production is able to enhance
XMP production
Acknowledgements
We are grateful to K Sato and T Kazarimoto for their helpful input.
References
1 Sauer, U & Bailey, J.E (1999) Estimation of P-to-O ratio in
Bacillus subtilis and its influence on maximum riboflavin yield.
Biotechnol Bioeng 64, 750–754.
2 Dauner, M & Sauer, U (2001) Stoichiometric growth model
for riboflavin-producing Bacillus subtilis Biotechnol Bioeng 76,
132–143.
3 Barrow, K.D., Collins, J.G., Norton, R.S., Rogers, P.L & Smith,
G.M (1984) 31 P nuclear magnetic resonance studies of the
fer-mentation of glcose to ethanol by Zymomonas mobilis J Biol.
Chem 259, 5711–5716.
4 Castro, C.D., Koretsky, A.P & Domach, M.M (1999)
Perfor-mance trade-offs in in vivo chemostat NMR Biotechnol Prog 15,
185–195.
5 Noguchi, Y., Shimba, N., Toyosaki, H., Ebisawa, K., Kawahara,
Y & Suzuki, Ei & Sugimoto, S (2002) In vivo NMR system for evaluating oxygen-dependent metabolic status in microbial culture J Microbiol Methods 51, 73–82.
6 Dulyaninova, N.G., Podlepa, E.M., Toulokhonova1, L.V & Bykhovsky, V.Y (2000) Salvage pathway for NAD biosynthesis
in Brevibacterium ammoniagenes: regulatory properties of triphos-phate-dependent nicotinate phosphoribosyltransferase Biochim Biophys Acta 1478, 211–220.
7 Han, J.K., Chung, S.O., Lee, J.H & Byun, S.M (1997) 6¢-Mercaptoguanosine-resistance is related with purF gene encoding 5¢-phosphoribosyl-1¢-pyrophosphate amidotransferase
in inosine-5¢-monophosphate overproducing Brevibacteirum ammoniagenes Biotechnol Lett 19, 79–83.
8 Usuda, Y., Kawasaki, H & Utagawa, T (2001) Characterization
of the cell surface protein gene of Corynebacterium ammoniagenes Biochim Biophys Acta 1522, 138–141.
9 Neves, A.R., Ramos, A., Nunes, M.C., Kleerebezem, M., Huge-nholtz, J., de Vos, W.M., Almeida, J & Santos, H (1999) In vivo nuclear magnetic resonance studies of glycolytic kinetics in Lactococcus lactis Biotechnol Bioeng 64, 200–212.
10 Crosse, A.M., Greenway, D.L & England, R.R (2000) Accu-mulation of ppGpp and ppGp in Staphylococcus aureus 8325–4 following nutrient starvation Lett Appl Microbiol 31, 332–337.
11 Meyer, S., Noisommit-Rizzi, N., Reuss, M & Neubauer, P (1999) Optimized analysis of intracellular adenosine and guanosine phosphates in Escherichia coli Anal Biochem 271, 43–52.
12 Lundberg, P., Harmsen, E., Ho, C & Vogel, H.J (1990) Nuclear magnetic resonance studies of cellular metabolism Anal Biochem.
191, 193–222.
13 Greiner, J.V., Kopp, S.J & Glonek, T (1985) Phosphorus nuclear magnetic resonance and ocular metabolism Surv Ophthalmol 30, 189–202.
14 Dauner, M., Bailey, J.E & Sauer, U (2001) Metabolic flux ana-lysis with a comprehensive isotopomer model in Bacillus subtilis Biotechnol Bioeng 76, 144–156.
15 Sauer, U., Hatzimanikatis, V., Hohmann, H.P., Manneberg, M., van Loon, A.P & Bailey, J.E (1996) Physiology and metabolic fluxes of wild-type and riboflavin-producing Bacillus subtilis Appl Environ Microbiol 62, 3687–3696.
16 Dauner, M., Sonderegger, M., Hochuli, M., Szyperski, T., Wuthrich, K., Hohmann, H.P., Sauer, U & Bailey, J.E (2002) Intracellular carbon fluxes in riboflavin-producing Bacillus subtilis during growth on two-carbon substrate mixtures Appl Environ Microbiol 68, 1760–1771.
17 Kovarova-Kovar, K., Gehlen, S., Kunze, A., Keller, T., Daniken, R.V., Kolb, M & van Loon, A.P (2000) Application of model-predictive control based on artificial neural networks to optimize the fed-batch process for riboflavin production J Biotechnol 79, 39–52.