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Journal of Bacteriology, February 2000, p. 620-626, Vol. 182, No. 3
0021-9193/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
Extracellular Oxidoreduction Potential Modifies
Carbon and Electron Flow in Escherichia coli
Christophe
Riondet,1,2
Rémy
Cachon,1,*
Yves
Waché,1
Gérard
Alcaraz,2 and
Charles
Diviès1
Laboratoire de Microbiologie U.A. INRA,
ENSBANA, Université de Bourgogne,1 and
Laboratoire de Physiologie et de Biochimie
Végétale, ENESAD,2 21000 Dijon,
France
Received 8 July 1999/Accepted 3 November 1999
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ABSTRACT |
Wild-type Escherichia coli K-12 ferments glucose to a
mixture of ethanol and acetic, lactic, formic, and succinic acids. In anoxic chemostat culture at four dilution rates and two different oxidoreduction potentials (ORP), this strain generated a spectrum of
products which depended on ORP. Whatever the dilution rate tested, in
low reducing conditions (
100 mV), the production of formate, acetate,
ethanol, and lactate was in molar proportions of approximately
2.5:1:1:0.3, and in high reducing conditions (320 mV), the production
was in molar proportions of 2:0.6:1:2. The modification of metabolic
fluxes was due to an ORP effect on the synthesis or stability of some
fermentation enzymes; thus, in high reducing conditions, lactate
dehydrogenase-specific activity increased by a factor of 3 to 6. Those
modifications were concomitant with a threefold decrease in
acetyl-coenzyme A (CoA) needed for biomass synthesis and a 0.5- to
5-fold decrease in formate flux. Calculations of carbon and cofactor
balances have shown that fermentation was balanced and that
extracellular ORP did not modify the oxidoreduction state of cofactors.
From this, it was concluded that extracellular ORP could regulate both
some specific enzyme activities and the acetyl-CoA needed for biomass
synthesis, which modifies metabolic fluxes and ATP yield, leading to
variation in biomass synthesis.
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INTRODUCTION |
A wealth of information is available
on the response of Escherichia coli cellular metabolism to
pH, water activity, or temperature variations, but little is known
about the action of extracellular oxidoreduction potentials (ORP) on
metabolism, although numerous reactions and regulations are of the
oxidoreduction type. Previous studies have shown that substrates with
different oxidation states yield a specific product spectrum. Thus,
with glucose (oxidation state = 0), the spectrum of main end
products (formate:acetate:ethanol:lactate) is equal to 2:1:1:2, with
glucitol (oxidation state = 1), it is equal to 2:1:6:0.5, and
with glucuronate (oxidation state = 2), it is equal to 1:5:1:1,
with small amounts of succinate also being produced in each case
(1). Those metabolic flux modifications are also observed
when the nature of external electron acceptors varies (7).
In the same way, the NADH/NAD+ ratio, which is responsible
for regulation of enzymes (8) or genes (16), can
be influenced by the oxidation level of the substrate (36)
or by the availability and nature of electron acceptors (7).
In addition, protein folding and disulfide bond formation are regulated
and modified by different oxidoreductase enzymes and oxidoreduction
couples (glutathione, thioredoxin) (26). Recently, Taylor
and Zhulin in their review have shown that ORP influences or could
influence numerous regulations of cell functions controlled via
Per-Arnt-Sim (PAS)-containing receptors (signaling modules that monitor
changes in light, ORP, oxygen, and the overall energy level of a cell),
transducers, and regulators (34). Thus, E. coli
senses the medium ORP and swims to a preferred ORP niche by redox
taxis, involving a change in proton motive force as a hypothetical
sensor (3).
Few experimental studies have explored the extracellular ORP effect on
particular metabolite production. Kwong and Rao have shown that
reducing conditions can increase productions of homoserine and lysine
by Clostridium glutamicum (13). In the same way, for Clostridium acetobutylicum, a decrease from 300 to
370 mV increases the production level of butanol and decreases those of butyrate and acetate (24). These results show that
extracellular ORP can modify metabolic fluxes.
To our knowledge, however, a more global study of ORP action on
metabolism has not been realized, although Gill et al. have shown that
extracellular ORP could act on the oxidoreduction state of
cytoplasmic molecules, due to the low ORP buffer strength of cytoplasm
(9). Furthermore, from a biotechnological point of view, if
this parameter can modify metabolic fluxes, it might be an additional
physicochemical parameter to take into account for the optimization of processes.
In this paper, we describe the effect of ORP on modifications of the
metabolic end product spectrum (lactate, acetate, ethanol, formate,
succinate, and CO2), enzyme activities (alcohol
dehydrogenase [ADH], lactate dehydrogenase [LDH], acetate kinase
[AK], pyruvate kinase [PK], and phosphoenolpyruvate carboxylase
[PEPC]), intermediary metabolites (acetyl-coenzyme A [CoA],
oxaloacetic acid [OAA], and phosphoenolpyruvate [PEP]),
oxidized and reduced forms of NAD in E. coli, and ORP
modified fermentation pathways, leading to changes in end product
spectrum, growth yield, and oxidation balance.
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MATERIALS AND METHODS |
Batch cultivation: bacterial strain and culture conditions.
E. coli K-12 wild-type EMG2 (CGSC 4401) was grown in
Trypticase soy broth sterilized by filtration, in anaerobic conditions. Cells were cultivated in an 800-ml bioreactor (Biostat Q, Braun, Germany). Temperature, pH, and agitation were maintained at 37°C, 7.0 (with NaOH as neutralizer), and 200 rpm respectively. ORP was measured
with a redox combined electrode (Pt 4805-DXK; Mettler-Toledo S.A.R.L.,
Paris, France) connected to a redox controller (P507 Consort; Bioblock
Scientific, Illkirch, France), and the values were corrected according
to the reference electrode value (210 mV at 37°C). Before
inoculation, the medium was sparged with nitrogen to eliminate oxygen.
Metabolic end products were measured at the beginning of the stationary phase.
Chemostat cultivation: culture conditions.
The strain was
grown in minimal M9 medium with glucose (20 g/liter) as the carbon
source, sterilized by filtration. An anaerobic cultivation condition
was chosen to avoid the uncertainties associated with the P/O ratio of
oxidative phosphorylation for ATP yield calculations. In anaerobic
growth without external electron acceptors such as nitrate or fumarate,
energy generation occurs only through substrate level phosphorylation.
Chemostat cultivation was chosen over batch cultivation to provide a
better quality of specific rate data.
The chemostat cultivation was performed in a 2.0-liter bioreactor
(Setric G.I., Toulouse, France) equipped with a weight control for
maintaining a constant working volume of 1.5 liters. After inoculation
of the fermentor, it was run in batch mode overnight and then put on
chemostat condition. Four dilution rates (0.025, 0.05, 0.1, and 0.2 h
1) were studied. In these conditions, glucose was not a
limiting nutrient. The continuous culture reached a steady state after 5 to 6 residence times. The pH, temperature, and agitation were maintained at 6.0 (with NaOH as a neutralizer), 37°C, and 200 rpm,
respectively. Two distinct sets of ORP conditions were maintained, the
first set, named low reducing conditions (LRC), was obtained by a
sparging nitrogen gas flow which dispelled the hydrogen produced naturally by the bacteria (mean ORP ± standard deviation,
90 ± 50 mV, depending on the dilution rate). The second set,
named high reducing conditions (HRC), was obtained by a low sparging nitrogen gas flow, with, if necessary, hydrogen added to maintain a
mean ORP of 320 mV (standard deviation, 20 mV, depending on the
dilution rate). To limit the end product evaporation loss (especially
ethanol) by gas flow, the fermentor was equipped with an exhaust gas
cooler, maintained at 0°C by a refrigerated circulating bath. ORP was
measured with a redox combined electrode. To ensure that all oxygen
traces were removed from the feeding medium, it was sparged with
oxygen-free nitrogen before use.
Analytical technique.
Cell density was monitored at 620 nm
in a spectrophotometer (Novaspec 4049; LKB Products) and reported as
cellular dry weight. For substrate and fermentation end product
measures, a 10-ml sample was centrifuged twice (20,000 × g, 10 min at 4°C) and the supernatant was frozen at 80°C
until analysis. Glucose, acetate, ethanol, succinate, formate, and
lactate were quantified using a high-pressure liquid chromatography
system (Shimadzu) equipped with a cation-exchange column (HPX-87H;
Bio-Rad Labs), a differential refractive index detector, and a UV
detector. A mobile phase pH of 1.32 (H2SO4) at a 0.5-ml/min flow rate was
used, and the column was operated at 60°C. CO2
production was measured by passing the effluent gas from the fermentor
through a CO2 analyzer (infrared absorption technology;
Leybold Heraeus).
For enzyme activity determination, a 200-ml sample was centrifuged
(6,000 ×
g, 10 min at 4°C), washed twice with 50 mM
potassium
phosphate buffer (pH 7.5), resuspended in an equal protein
volume
at a concentration of 10 g · liter
1 and
frozen for no more than 2 days at
20°C (
15). To obtain
the cell extracts, cells were sonicated three times for 3 min
using
cycles of 1 s of sonication (power 7) and 1 s of rest (Vibra
cell disruptor; Sonics Materials, Danburry, Conn.) and cooled
in an
ice-water bath. The cell debris was removed by centrifugation
(20,000 ×
g, 10 min at 4°C). The supernatant was
used for the
measurements of the activities of the five enzymes. All
enzyme
assays were realized in the linear response range (determined
beforehand) of each enzyme by adequately diluting the
sample.
LDH was assayed by the method described by Racker (
25), in a
reaction mixture containing potassium phosphate buffer (100
mM, pH
7.5), NADH (0.33 mM), and sodium pyruvate (30 mM). The
reaction was
initiated by the addition of
pyruvate.
AK was assayed by the method described by Nakajima et al.
(
21), in a reaction mixture consisting of imidazole-HCl
buffer
(50 mM, pH 7.3), MgCl
2 (10 mM), glucose (10 mM),
NADP (1.6 mM),
acetyl phosphate (12 mM), ADP (5 mM), hexokinase (56 U/ml), and
glucose 6-phosphate-dehydrogenase (1.5 U/ml). The reaction
was
initiated by the addition of
ADP.
ADH was assayed by the method described by Clark and Cronan, Jr.
(
5), in a reaction mixture containing sodium phosphate
buffer (12 mM, pH 8.5), ethanol (300 mM), and NAD (0.3 mM). The
reaction was initiated by the addition of
ethanol.
PEPC was assayed by the method described by McAlister et al.
(
18), in a reaction mixture containing Tris-HCl buffer (100
mM, pH 8.5), PEP (6 mM), MgSO
4 (5 mM), KHCO
3
(10 mM), acetyl-CoA
(1 mM), NADH (0.3 mM), and malate dehydrogenase (10 U/ml). The
reaction was initiated by the addition of acetyl-CoA.
PK was assayed by the method described by Garrigues et al.
(
8), in a reaction mixture containing Tris-HCl buffer (100 mM,
pH 7.2), MnSO
4 (5 mM), KCl (10 mM), ADP, (1 mM), NADH
(0.3 mM),
LDH (10 U/ml), and PEP (2 mM). The reaction was initiated by
the
addition of
PEP.
Protein concentration was determined by the method of Lowry et al.
(
17); bovine serum albumin was used as the standard.
One
unit of enzyme is defined as the amount of enzyme required
to produce 1 µmol of product per
min.
In vitro activity and stability of LDH have been measured in the two
sets of ORP conditions, LRC and HRC, generated by a sparging
gas flow
of nitrogen and hydrogen, respectively, which created
ORP conditions as
close as possible to those utilized during batch
cultivation. Cell
extracts containing LDH were obtained from a
batch cultivation of
E. coli in M9 medium harvested at a culture
pH of 6.0 and
lysed as described above. LDH activity was measured
with the LDH assay
described above. In the case of in vitro LDH
activity measurement, the
two ORP conditions were generated with
a gas sparging into the
different solutions and maintained during
the assay with a continued
gas flow in the spectrometer cuvette.
In the case of the in vitro LDH
stability measurement (pH 7.5,
50°C), the two sets ORP conditions
were applied to the cell extracts
with gas sparging before and during
heat treatment, and then LDH
activity was
determined.
Intracellular metabolite and coenzyme concentrations were measured by
using an in vitro procedure based on rapid inactivation
of metabolism
followed by metabolite extraction directly from
the cell sample. Cell
samples were removed from the culture, and
metabolite extraction was
performed immediately. Methods of metabolite
extraction (acidic or
basic) and assays were based on those developed
by Lebloas
(
15). A variable volume of either HCl (6 N) or KOH
(10 N)
was added to yield a final pH of 1.2 or 12.5, respectively.
Most
metabolites and coenzymes were extracted by incubating the
HCl-treated
culture at 50°C for 10 min before neutralizing it
to a pH of 6.5 to 7 by KOH during vigorous agitation. After a
10-min centrifugation
(6,000 ×
g at 4°C), the supernatant was
immediately
used for metabolite concentration measurements. Acid-labile
coenzymes,
like NADH, were extracted by incubating the KOH-treated
culture (pH
12.5) for 10 min at room temperature (25°C). After
centrifugation
(6,000 ×
g, 10 min at 4°C), the supernatant was
immediately tested for NADH. Metabolites were measured by coupling
appropriate enzyme assays with fluorimetric determination of the
coenzyme NADH or NADPH. Emission was measured at 460 nm after
excitation at 350 nm with a spectrofluorometer (Hitachi F-4500;
Hitachi
Instrument Co., Tokyo,
Japan).
Pyruvate was assayed in a reaction mixture containing potassium
phosphate buffer (100 mM, pH 7), MgCl
2 (2.5 mM), NADH (33
µM), acid extract, and LDH (20 U · ml
1) to
initiate pyruvate consumption (
6).
PEP was determined after complete depletion of the pyruvate present in
the extract by using a mixture which contained potassium
phosphate
buffer (100 mM, pH 7), MgCl
2 (2.5 mM), ADP (2 mM), a
molar
concentration of NADH sufficient to complete pyruvate elimination,
acid
extract, and LDH (20 U · ml
1). After all the
pyruvate had been consumed, the level of remaining
NADH was checked and
adjusted to obtain the PEP measure and pyruvate
kinase (5 U · ml
1) was added to initiate PEP assay (
6).
Acetyl-CoA was assayed in a reaction mixture containing Tris-HCl buffer
(200 mM, pH 8.1), malate (5 mM), NAD
+ (33 µM), acid
extract (first reading), malate dehydrogenase (1
U · ml
1) (second reading), and citrate synthase (0.08 U
· ml
1) (third reading) (
20).
Oxaloacetate was assayed in a reaction mixture containing potassium
phosphate buffer (100 mM, pH 7), MgCl
2 (2.5 mM), NADH
(33 µM), acid extract, and MDH (120 U · ml
1) to
initiate oxaloacetate consumption (procedure was adapted
from that
described in reference
18).
NAD
+ was assayed in a reaction mixture containing
pyrophosphate buffer (50 mM, pH 8.8), semicarbazide (2.5 g · liter
1), absolute ethanol (80 mM), acid extract, and
alcohol dehydrogenase
(0.5 U · ml
1)
(
12). The NADH present in the alkali extract was measured
in
a reaction mixture containing triethanolamine buffer (200 mM,
pH 7),
pyruvate (5 mM), alkaline extract, and LDH (20 U · ml
1) to initiate the pyruvate-dependent oxidation of NADH
(
11).
All metabolite concentrations obtained were relative to cellular dry
weight but were expressed as aqueous molar values, using
an average
intracellular concentration of 2.3 ml · g (dry
weight)
1 (
10).
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RESULTS |
Batch cultivation.
The carbon recovery levels were close to
97% in the three experiments (Table 1).
When the ORP increased from 360 to 280 mV, lactate production
decreased from 0.15 to 0.01 mol per mol of consumed substrate, carbon
dioxide decreased from 0.71 to 0.39 mol per mol of consumed substrate,
and formate increased from 0.88 to 1.31 mol per mol of consumed
substrate. In fact, the total formate production, equal to the sum of
ethanol and acetate (Fig. 1), did not
vary with ORP values, but CO2 produced from formate decreased when the ORP increased. The three other products, acetate, ethanol, and succinate, did not seem to be affected by ORP variations. Concerning the final biomass, the fermentation with 1 g of
dithiothreitol (DTT) per liter presented a decrease of 30% compared to
normal growth. The weight of dry biomass produced per ATP mole
(YATP) varied from 5.6 to 7.9 g (dry
weight) · mol of ATP1.
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TABLE 1.
Effect of ORP on metabolic end product distribution for
anaerobic batch cultivation in Trypticase soy broth medium
maintained at a pH of 7.0
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Chemostat cultivation. (i) Effect of the ORP on growth.
The
biomass was affected by the ORP value and by the dilution rate (µ).
When the µ increased from 0.025 to 0.2 h1, in LRC,
biomass decreased from 0.49 to 0.20 g · liter1,
and in HRC biomass decreased from 0.2 to 0.12 g · liter1. Whatever the tested dilution rate was, a 250-mV
ORP decrease led to a twofold decrease in biomass production
(Table 2). The maximum
YATP
(YATPmax) and
maintenance coefficient (mATP, in mmoles of
ATP · g [dry weight]1 · h1)
values were identical under the two sets of ORP conditions (Fig. 2). The glucose consumption was on
average two times higher in LRC than in HRC (Table 2). As for the
average ATP yield, there was a slight decrease from 2.8 mol of ATP
· mol of consumed glucose1 in LRC to 2.3 mol of
ATP · mol of consumed glucose1 in HRC (Table
2).
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TABLE 2.
Results of anaerobic chemostat cultivation in minimum
media with glucose as the carbon source maintained at a pH of 6.0
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FIG. 2.
Effect of the dilution rate and ORP conditions (LRC
[filled symbols] and HRC [open symbols]) on
YATP of E. coli K-12 when it was
grown in continuous culture.
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(ii) Metabolic fluxes and balances.
The carbon recovery level
was between 90 and 98%, measured independently of the dilution rate
and ORP conditions (Table 2). The distribution of the consumed glucose
carbon between the different products of glucose metabolism was
calculated for the eight conditions (Fig.
3). The elemental composition
C4.2H8O1.25N0.68P0.1
was used to estimate the amount of assimilated carbon recovered in
biomass (22). In the case of LRC, the product distribution,
expressed in moles per mole of consumed substrate, was identical for
all dilution rates. The fermentation produced formate (formate and CO2), acetate, ethanol, and lactate in proportions of
approximately 2.5:1:1:0.3 and a small amount of succinate and
CO2. With the HRC, this proportion was radically modified
by increased production of lactate and CO2 and decreased
production of acetate (Fig. 3), and the fermentation spectrum was on
average 2:0.6:1:2, with a small amount of succinate. The lactate
percentage increased from 10 to 12% of the total carbon in LRC to 35 to 47% of the total carbon in HRC. As for the ethanol/acetate ratio,
it increased from 1.07 to 1.15 in LRC to 1.33 to 1.42 in HRC. The total
carbon dioxide synthesis (CO2 gas measured and the part
used in succinate synthesis) was about 2% in LRC and between 3.5 and
9% (depending on the dilution rate) in HRC. The NADH/NAD+
ratio was independent of ORP conditions but varied with the dilution rate from 0.2 in HRC to 0.5 in LRC (Fig.
4). The NADH recovery, was on average
equal to 115%, independent of the dilution rate and ORP (Table 2).
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FIG. 3.
Production rate of formate, acetate, ethanol, lactate,
succinate, and CO2 in anaerobic glucose chemostat culture
of E. coli K-12 regulated at a pH of 6.0, depending on
dilution rate, and in LRC (range, 40 to 140 mV) (A) and HRC (range,
315 to 340 mV) (B). Each value is the average of two measurements.
The standard deviation is ±10%.
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FIG. 4.
NADH/NAD+ ratio versus dilution rate and ORP
conditions (LRC [black] and HRC [gray]). The values are the
means ± standard deviations (error bars) of at least three
measurements.
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(iii) Oxidation balance.
Each molecule has its own oxidation
state (glucose, 0; acetate, 0; lactate, 0; formate, +1; succinate, +1;
carbon dioxide, +2; hydrogen, 1; and ethanol, 2). Using the average
composition formula of E. coli,
C4.2H8O1.25N0.68P0.1
and 4% for trace elements (
100 g/mol) (22), the
oxidation state of the biomass is equal to 1.5. The sum of oxidized
and reduced products should equal the oxidation state of the substrate
(4). Whereas for HRC, there was an excess of reducing
equivalents of 0.12 to 0.27 mol per mol of glucose, for LRC, the excess
of oxidized equivalents was between 0.30 and 0.64 mol per mol of
glucose (Table 2).
(iv) Enzyme synthesis.
The activities of the PEPC, ADH, and PK
were constant whatever the dilution rate, whereas the activity of LDH
decreased and that of AK increased with an increase in the dilution
rate, for the two sets of ORP conditions (Fig.
5). As for ORP, it especially affected
the LDH and PEPC activities. The former increased three- to sixfold and
the latter increased two- to fivefold in HRC compared to LRC.
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FIG. 5.
Effect of dilution rate and the two sets of ORP
conditions (LRC and HRC) on in vitro measurements of LDH, AK, ADH,
PEPC, and PK in E. coli K-12 cell extract. For each enzyme
100% was equal to the highest values obtained whatever the dilution
rate and ORP condition. Each value is the average ± standard
deviation (error bar) of at least three measurements.
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(v) Intracellular intermediary metabolites.
PEP,
acetyl-CoA, OAA, and pyruvate are situated at four major branch
points of glycolysis and fermentation pathways which intervene in
numerous enzyme regulations. Due to a high extracellular concentration
of pyruvate, the difference between intra- and extracytoplasmic concentrations was less than the assay error, and consequently, the
intracellular concentration of pyruvate was not determined. But it was
interesting to note that the extracellular concentration of pyruvate
was on average 10-fold higher in HRC than in LRC (Table 3).
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TABLE 3.
Intracellular concentrations of three intermediary
metabolites and of extracellular pyruvate, depending on dilution
rate and ORP conditionsa
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PEP, acetyl-CoA, and OAA concentrations increased five to tenfold when
the dilution rate increased from 0.025 to 0.2 h
1.
Concerning ORP effect, there was no significant difference for
PEP and
OAA, but acetyl-CoA concentration increased two- to eightfold
in HRC
(Table
3).
(vi) Steady-state fluxes of end products.
The glucose
flux (qglucose) slightly increased in HRC
(Table 4). In the cases of acetate flux
(qacetate), ethanol flux
(qethanol), and succinate flux
(qsuccinate), the ORP had no significant effect. On the other hand, in HRC, lactate flux
(qlactate) increased three- to sixfold, the
CO2 flux (qCO2) increased 8- to
10-fold and the formate flux (qformate)
decreased from 0.5- to 5-fold. Thus, pyruvate was preferentially used
for lactate synthesis in HRC and for acetyl-CoA and formate production
in LRC. The difference between total formate flux
(qformate plus qCO2 plus
qsuccinate) and the sum of
qethanol and qacetate
showed that acetyl-CoA and derived compounds were used for biomass
synthesis. This need varied on average from 38% of total acetyl-CoA
flux in LRC to 12% of total acetyl-CoA flux in LRC.
(vii) In vitro activity and stability of LDH.
In vitro
activity and stability at 50°C of LDH were identical in the two ORP
conditions tested (Fig. 6).
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FIG. 6.
Effect of ORP (LRC [filled symbols] and HRC [open
symbols]) on in vitro activity (A) and stability (B) of LDH. Activity
of 100% corresponds to the activity of LDH at a pH of 7.5 in LRC. The
values are the averages ± standard deviations of two independent
measurements.
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DISCUSSION |
The study realized in batch culture shows that ORP affects
metabolic fluxes. The main results were a decrease in lactate and CO2 production and an increase in formate production when
the ORP increased (Table 1). These results were obtained in batch culture using DTT and sodium borohydride to modify the ORP.
Because of the known toxicity of DTT to E. coli
(2) and because of the instability of sodium borohydride in
solution, another reducing compound, hydrogen, recognized for its
absence of toxicity, has been chosen for further experiments. The
results obtained with chemostat cultivation confirm results obtained in
the preliminary study and show clearly that changes in ORP
affect metabolic fluxes of E. coli, but without modification
of YATPmax and
mATP values.
In HRC, a decrease in the level of biomass synthesis was observed,
since ATP yield decreased from 2.8 to 2.3 mol of ATP · mol of
consumed glucose1 and substrate consumption was divided
by two (Table 2). When ORP decreased, lactate production increased
essentially to the detriment of formate and in a lower proportion of
acetate. Thus, for an average ORP of 100 mV, 10% of carbon was
transformed into lactate, and this percentage increased to 34 to 46%
for an average ORP of 320 mV, for all tested dilution rates.
Furthermore, for HRC and LRC respectively, acetyl-CoA used for biomass
synthesis represented between 11 and 36% of acetyl-CoA flux. Decreases
in acetyl-CoA need and in formate flux resulted in acetyl-CoA and pyruvate accumulation.
Measurements in HRC of the specific activities of LDH were in agreement
with those of lactate flux, increasing by a factor 3.5 to 6 for LDH
activity (Fig. 5) and 3.5 to 5.0 for lactate flux (Table 4), compared
to values obtained in LRC. The similarity of variation between
LDH-specific activity and lactate production shows that the apparent
activity of LDH is independent of extracellular ORP conditions.
Concerning the LDH activity, it could be assumed that its activity was
not ORP sensitive, based on the knowledge that E. coli
cytoplasm is not well ORP buffered, as has been demonstrated already
(9). Measurement of in vitro activity of LDH confirmed the
assumption (Fig. 6). To our knowledge, only a few enzyme
activities of E. coli are classified as sensitive
to ORP: the phosphotransferase system (PTS) (30), the
sugar-proton and amino acid-proton symports (12, 29), and,
as was more recently demonstrated, the glutathione-gated K+
channels (19).
Thus, modifications of metabolic fluxes are linked to variations of
specific enzyme activities, principally an increase of LDH and probably
of formate dehydrogenase (FDH), as observed by the carbon dioxide
production in HRC, which was six to seven times higher than that in LRC
(Fig. 3). These two enzymes are known to be overexpressed in acidic
conditions (4), but chemostat is pH regulated and the higher
activity observed cannot be due to extracellular pH variations.
However, we have shown recently that on resting cells of E. coli, reducing conditions permeabilized the cytoplasmic membrane
mainly to protons (28).
It should thereby be assumed that if the cytoplasmic membrane is
permeabilized, the intracellular pH is lower than in less reducing
conditions. The cell has different methods for detecting internal and
external pH (27, 33), but the mechanisms of internal pH
detection are not clearly understood. Although to our knowledge only a
few genes show pH-dependent induction or overexpression, like the
multiple drug resistance regulon mar (31), the
ferric iron uptake regulation gene fur, and others
identified or not (14), the regulation of other genes could
be sensitive like LDH, and perhaps FDH, to an acidification of
cytoplasm without external pH variation, as our results seem to show.
Another explanation may be that the enzyme has an ORP-dependent
expression and/or protein stability. This last hypothesis has been
tested, but no difference between results for LRC and HRC could be
shown (Fig. 6). An ORP-dependent expression, as has been already
demonstrated for the fumarate nitrate reductase (FNR) system
(35) and for the adenosyl-cobalamin biosynthetic (Ado-CBL) operon responsible for coenzyme B12 synthesis
(23), seems to be the right hypothesis to explain this
modification of LDH-specific activity.
Concerning the ethanol/acetate ratio, in HRC, on average it was equal
to 1.36, and it decreased to 1.12 at 100 mV. On the basis of NADH
recovery, a normal ratio would be equal to 1. It has been shown that
hydrogen produced by the formate hydrogenlyase (FHL) complex could be
recycled by the hydrogenases Hyd-1 and/or Hyd-2 in fermentation
conditions to increase the reductive pathway with ethanol formation
(1, 32). This is what we observed in our study, and due to
the nearly identical NADH recovery in the two ORP conditions, it can be
supposed that extra reducing equivalents are directly transferred to
ethanol synthesis, which slightly reduces acetate production in HRC.
The redox balance, which presents a molar excess of 0.12 to 0.27 mol of
reducing equivalents per mol of consumed glucose, tends to demonstrate this. On the contrary, in LRC, the redox balance presents an excess of
oxidant equivalents (0.30 to 0.64 mol of oxidant equivalents per mol of
consumed glucose), which confirms that extracellular ORP can modify the
oxidation balance of metabolism.
The NAD cofactor ratio and NADH recovery are not modified by
extracellular ORP, which illustrates the necessity of a balance in the
rates of oxidation and reduction of these nucleotides to ensure the
continuation of both catabolism and anabolism for a given carbon source
in E. coli.
From this study, it can be concluded that manipulating the ORP changes
the proportion of metabolic end products. The two assumed main causes
involved in such a shift are alterations of the specific enzyme
activities of LDH and FDH and of the acetyl-CoA requirement for biomass synthesis.
In fact, ORP modifies the biomass synthesis, affecting the ratio of
glucose consumed while the YATPmax
and the mATP values remain constant. Such a
modification might be explained by an overall activation or inhibition
of the microbial physiology in different oxidoreduction conditions.
| |
ACKNOWLEDGMENTS |
This work was supported by the program Aliment Demain contract
R97/04 of the Ministère de l'Agriculture, de la Pêche et de l'Alimentation. C. Riondet holds doctorate fellowships from the Conseil Régional de Bourgogne.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Laboratoire de
Microbiologie, A.U. INRA, ENSBANA, 1 esplanade Erasme, 21000 Dijon, France. Phone: (33) 03 80 39 66 73. Fax: (33) 03 80 39 66 40. E-mail:
cachon{at}u-bourgogne.fr.
| |
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Abstract
Introduction
