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Applied and Environmental Microbiology, July 2000, p. 2934-2942, Vol. 66, No. 7
0099-2240/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
Energetics of Syntrophic Propionate Oxidation in
Defined Batch and Chemostat Cocultures
Johannes C. M.
Scholten
and
Ralf
Conrad*
Max-Planck-Institut für terrestrische
Mikrobiologie, D-35043 Marburg, Germany
Received 9 February 2000/Accepted 8 May 2000
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ABSTRACT |
Propionate consumption was studied in syntrophic batch and
chemostat cocultures of Syntrophobacter fumaroxidans and
Methanospirillum hungatei. The Gibbs free energy available
for the H2-consuming methanogens was <
20 kJ mol of
CH4
1 and thus allowed the synthesis of 1/3
mol of ATP per reaction. The Gibbs free energy available for the
propionate oxidizer, on the other hand, was usually >10 kJ mol of
propionate1. Nevertheless, the syntrophic coculture grew
in the chemostat at steady-state rates of 0.04 to 0.07 day1 and produced maximum biomass yields of 2.6 g
mol of propionate1 and 7.6 g mol of
CH41 for S. fumaroxidans and
M. hungatei, respectively. The energy efficiency for
syntrophic growth of S. fumaroxidans, i.e., the biomass
produced per unit of available Gibbs free energy was comparable to a
theoretical growth yield of 5 to 12 g mol of ATP1.
However, a lower growth efficiency was observed when sulfate served as
an additional electron acceptor, suggesting inefficient energy
conservation in the presence of sulfate. The maintenance Gibbs free
energy determined from the maintenance coefficient of syntrophically
grown S. fumaroxidans was surprisingly low (0.14 kJ
h1 mol of biomass C1) compared to the
theoretical value. On the other hand, the Gibbs free-energy dissipation
per mole of biomass C produced was much higher than expected. We
conclude that the small Gibbs free energy available in many
methanogenic environments is sufficient for syntrophic propionate
oxidizers to survive on a Gibbs free energy that is much lower than
that theoretically predicted.
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INTRODUCTION |
Propionate is an important
intermediate in the conversion of organic matter to methane and carbon
dioxide. In methanogenic environments, the degradation of propionate to
acetate and CO2 may account for 6 to 35 mol% of the total
methanogenesis (17). Propionate oxidation itself is
energetically very unfavorable under standard thermodynamic conditions
(see Table 1). Under methanogenic conditions, proton-reducing
acetogenic bacteria are only able to gain energy from this
reaction when the concentration of products is kept low. Thus, the
degradation of propionate is only accomplished in obligate syntrophic
consortia of proton-reducing acetogenic bacteria and methanogenic
archaea (27, 28, 33). So far, three syntrophic
propionate-oxidizing bacteria and some highly purified enrichment
cultures have been described (3, 11, 20, 21, 22, 32, 43, 44,
49).
Studies have shown that most of the known syntrophic
propionate-oxidizing bacteria degrade propionate via the
methylmalonyl-coenzyme A (CoA) pathway (14, 25, 43). During
the oxidation of propionate in the methylmalonyl-CoA pathway, electrons
are released in three reactions, namely, the oxidation of succinate to
fumarate, malate to oxaloacetate, and pyruvate to acetyl-CoA
(25). In methanogenic environments, the H2
partial pressure is low enough to allow the direct reduction of protons
with the electrons released during the oxidation of pyruvate and
malate. However, the H2 partial pressure is not sufficient
to allow this reduction during the oxidation of succinate to fumarate.
It was hypothesized that the electrons released during the oxidation of
succinate are shifted to a lower redox potential via reversed electron
transport. This transport would be driven by the hydrolysis of 2/3 mol
of ATP (27, 28, 37). Some evidence has been obtained for the
presence of a reversed electron transport system in syntrophic
propionate-degrading bacteria (41). However, the
methylmalonyl-CoA pathway yields only 1 mol of ATP via substrate level
phosphorylation. Therefore, if such a reversed electron transport is
occurring, only 1/3 mol of ATP per mol of propionate is left for
growth. Under physiological conditions, the Gibbs free-energy change
needed for ATP synthesis must amount to a minimum of 70 kJ mol of
ATP1. Thus, the minimum Gibbs free-energy quantum that
can generate 1/3 mol of ATP would amount to approximately 23 kJ mol
of propionate1. It has been suggested that this amount of
Gibbs free-energy change corresponds to the minimum energy quantum
required to sustain microbial life (27, 28).
In several methanogenic environments, the apparent Gibbs free-energy
change for propionate oxidation was on the order of 3 to 15 kJ mol
of propionate1 and thus was rather small (5, 19, 26,
46). This amount of free-energy change is less than the minimum
energy quantum needed to sustain microbial life. It is not clear why
such small free-energy changes are observed during the degradation of
propionate. Most syntrophic propionate-oxidizing bacteria are not
obligate syntrophs but are also able to grow on other substrates, such as fumarate, malate, and pyruvate, in the absence of a partner microorganism. A remarkable feature of the propionate-oxidizing Syntrophobacter species is their ability to couple the
oxidation of propionate not only to an H2-consuming
syntrophic partner but also to the reduction of sulfate. In fact,
phylogenetic analysis of Syntrophobacter fumaroxidans has
revealed that this bacterium is indeed related to sulfate-reducing
bacteria (12). Perhaps the ability to reduce sulfate is of
importance to explain the energetics of syntrophic propionate-oxidizing bacteria.
Besides propionate, many alcohols, fatty acids, amino acids, and
aromatic compounds are anaerobically degraded by syntrophy. In each
case, the available free energy is relatively low and has to be shared
between the two syntrophic partners (27, 28). The energetics
of syntrophic interspecies H2 transfer has been studied in
defined cocultures of benzoate-, lactate-, ethanol-, propionate-, and
butyrate-oxidizing fermenting bacteria with H2-consuming methanogens (1, 8, 9, 30, 31, 28, 39, 45). Propionate
oxidation, however, has not yet been investigated in continuous-culture experiments.
Therefore, we studied the energetics of propionate consumption in
syntrophic cocultures of S. fumaroxidans and
Methanospirillum hungatei in the absence and presence of
sulfate by determining the Gibbs free energy available for both the
propionate oxidizers and the H2-consuming methanogens under
steady-state conditions in batch and chemostat cultures. The growth
yields and maintenance coefficients of the syntrophic propionate
oxidizer were also determined.
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MATERIALS AND METHODS |
Organisms and cultivation.
M. hungatei
JF1T (DSM 864) and S. fumaroxidans MPOB (DSM
10017) were purchased from the Deutsche Sammlung von Mikroorganismen und Zellkulturen (Braunschweig, Germany).
The microorganisms were grown in bicarbonate-buffered, sulfide-reduced
mineral medium as described previously by Huser et al. (15).
To 1 liter of medium, 1 ml of a vitamin solution (48) and 1 ml each of an acid and an alkaline trace elements solution were added
(32). The vitamin solution was filter sterilized separately.
The gas phase above the medium was N2-CO2 (80%
to 20%) at 172 kPa, and the pH of the medium was 6.8 to 6.9. Substrates and other supplements were added from sterile anaerobic
stock solutions. Pure cultures of S. fumaroxidans and
M. hungatei were maintained on fumarate (40 mM) and
hydrogen, respectively. Hydrogen was added to the headspace at an
overpressure of 60 kPa. In the case of M. hungatei, 2 mM
(each) formate, acetate, and isobutyrate were added as supplementary
carbon sources. Culture purity was routinely checked by phase-contrast
and fluorescence microscopy.
Batch and chemostat experiments.
In batch experiments, pure
cultures of S. fumaroxidans or defined cocultures of
S. fumaroxidans and M. hungatei were cultivated in 1-liter serum bottles containing 500 ml of mineral medium with 40 mM
propionate with or without sulfate (40 mM) under a gas phase of
N2-CO2 (80% to 20%) at 172 kPa.
Continuous cultivation was performed in a 1-liter chemostat system as
described by Cypionka (
7) with a working volume of
800 ml
(flushed headspace) or 980 ml (nonflushed headspace). The
chemostat
experiments were only performed with cocultures of
S. fumaroxidans and
M. hungatei. The cocultures were grown
under
propionate limitation, and steady-state conditions were
maintained
for at least three culture volume changes. Substrate and
product
conditions were monitored, and total cell mass and species
composition
were checked under steady-state
conditions.
For inoculation of the batch and chemostat experiments,
S. fumaroxidans and
M. hungatei were pregrown on fumarate
and hydrogen,
respectively. Batch experiments with pure cultures of
S. fumaroxidans were inoculated with 10% (vol/vol)
S. fumaroxidans. Cocultures
for batch and chemostat
experiments were constructed by inoculating
10% (vol/vol)
S. fumaroxidans and 10% (vol/vol)
M. hungatei. All
experiments were performed at 37°C.
Determination of specific cell mass.
Batch experiments were
done as described above. Pure cultures of S. fumaroxidans
and M. hungatei were grown with propionate plus
SO42 and H2, respectively. Growth
was measured by monitoring the total cell protein (see below). Cells
were counted by phase-contrast microscopy using a Helber counting
chamber. Bacterial dry mass was determined gravimetrically
(see below). Cell suspensions contained (per liter) 27.4 ± 1.8 and 27.5 ± 0.6 mg (dry weight [dw]) of cells of S. fumaroxidans and M. hungatei, respectively. The
corresponding cell numbers were (108 per ml) 2.25 ± 0.81 and 2.33 ± 0.01. The corresponding specific cell masses
(xi; picograms [dw] per cell) 0.116 ± 0.037 and 0.119 ± 0.003.
Determination of growth parameters.
The maximum specific
growth rate (µmax; per day) was calculated from the
exponential part of the propionate depletion curves of batch cultures.
The total growth yield of the pure and mixed cultures growing on
propionate or on propionate plus sulfate in batch culture experiments
was determined from the microbial cell dw (see below). The total growth
yield in continuous culture was determined from the protein
concentration at steady state by multiplication by a conversion factor
(see below). The total biomass yield was calculated from the number of
grams of dry biomass produced per mole of propionate degraded. For
cocultures, the determined yield is the sum of the yields of both
species participating in the degradation of propionate. Consequently,
the measured yield is referred to as YXtot-S
(grams of dw per mole of a substrate).
The dry mass of the propionate-degrading and methanogenic
microorganisms (
Xi; grams [dw] per liter) were
calculated from the
total cell mass (
Xtot), the
relative cell numbers (
Ni; cells per
milliliter)
in the culture, and the specific cell masses
(
xi;
grams [dw] per cell) according to the
following equations (
26):
|
(1)
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(2)
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(3)
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The growth yields of the propionate-degrading microorganisms
(
YMPOB-S) and hydrogenotrophic methanogens
(
YMhun-P) in the
batch incubations were
calculated by relating the increase in
the biomass concentration to the
amount of propionate (S) degraded
or methane (P)
formed.
In the chemostat experiments, the maximum growth yield
(
YMPOBmax) and the maintenance
coefficient (
ms; millimoles of propionate
per
hour per gram [dw]) were obtained from the regression parameters
of
the following relationship (
24):
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(4)
|
with
Y the apparent growth yield at different
dilution rates (µ =
D) in a
chemostat.
Gibbs free-energy changes.
Standard Gibbs free-energy
changes (
G0) for the individual steps in the
degradation of propionate (see Table 1) were calculated from the
standard Gibbs free energies of formation
(Gf0) of the reactants and
products (36), corrected for a temperature of 37°C by the
Gibbs-Helmholtz equation. The Gibbs free energies (G) in
the cultures were calculated from the G0 of
the individual reactions using the actually measured concentrations or
partial pressures of the reactants and products, as well as the
prevailing temperature and pH (6.9).
Maintenance energy.
The maintenance energy
(mE) was determined from measured values as
described by Heijnen and VanDijken (13) and Tijhuis et al.
(38). The measured maintenance coefficients
(ms), in millimoles of substrate (electron
donor) per gram (dw) of biomass per hour, were converted to
mD, in moles of substrate C per mole of biomass C per hour. The available standard Gibbs free energy
(Gav01; kilojoules per mole of
substrate) was calculated from G0'
(kilojoules per mole of substrate) of the reaction divided by the
number of C atoms (three for propionate) and the degree of reduction
(
D = 4.67 for propionate) of the substrate. The
maintenance energy, in kilojoules per mole of biomass C per hour, was
then calculated by the following equation:
|
(5)
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Tijhuis et al. (
38) found that
mE can also be determined theoretically by the
following equation:
![m<SUB><UP>E</UP></SUB><UP> = 3.3 × exp</UP>[<UP>−6.94 × 10<SUP>4</SUP>/</UP>R(1/T − 1/298)]](/content/vol66/issue7/fulltext/2934/img006.gif)
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(6)
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Gibbs free-energy dissipation per mole of biomass C.
The
Gibbs free-energy dissipation per mole of biomass C
(Ds01/rAx;
kilojoules per mole of biomass C) was calculated as described by
Heijnen and VanDijken (13) by solving the macrochemical
equation, which, for the syntrophic oxidation of propionate by a
coculture of S. fumaroxidans and M. hungatei, can
be written as follows:
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(7)
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The macrochemical equation for the oxidation of propionate by
S. fumaroxidans can be written as follows:
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(8)
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The coefficient
f was calculated from the determined
maximum growth yield (
Ymax). The other five
stoichiometric coefficients (
a,
b,
c,
d,
e)
were
calculated from the five (C, H, O, N, and electric charge)
conservation
equations.
Ds01/
rAx was
then calculated by using the tabulated
Gf0 for the reactants and
products and
Gf0 =
67 kJ
mol of biomass C
1
(CH
1.8O
0.5N
0.2) (
13).
Heijnen and VanDijken (
13) showed that
Ds01/
rAx can
also be theoretically predicted from the number of C atoms
(
C) and the
degree of reduction (
yD)
of the substrate C source as follows:
![D<SUB><UP>sv</UP></SUB><SUP><UP>01</UP></SUP><UP>/</UP>r<SUB><UP>Ax</UP></SUB><UP> = 200 + 18</UP>(<UP>6 − </UP>C)<SUP>1.8</SUP> + <UP>exp</UP>{[(3.8 − y<SUB>D</SUB>)<SUP>2</SUP>]<SUP>0.16</SUP>(3.6 + 0.4C)}](/content/vol66/issue7/fulltext/2934/img011.gif)
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(9)
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For propionate, the theoretical
Ds01/
rAx is
426.6 kJ mol of biomass C
1. This value can then be used
to calculate a theoretical growth
yield for the coculture of
S. fumaroxidans plus
M. hungatei and
for
S. fumaroxidans alone using the macrochemical equations and
solving
for the six stoichiometric coefficients by using the six
(C, H, O, N,
electric charge, and energy) conservation equations
(
13).
Analytical procedures.
During microbial growth, samples were
taken for analysis of substrate and product concentrations.
H2 and CH4 were quantified by gas
chromatography (6, 29). Propionate and acetate were measured
by high-pressure liquid chromatography (18). Sulfate was
analyzed by ion chromatography (2). Bacterial growth was monitored by protein determination. Cell pellets of 4-ml culture samples were resuspended in 1 ml of 1 M NaOH. After heating at 100°C
for 15 min, the samples were treated further by the method of Bradford
(4). Bovine serum albumin was used as the standard. The dw
of a known culture volume was determined gravimetrically. The samples
were gassed with N2-CO2 to remove
H2S, centrifuged, and washed twice with 50 mM ammonium
acetate buffer (pH 6.0). The washed suspension was transferred into
glass bottles and dried at 100°C to a constant weight. Cell numbers
were counted by phase-contrast microscopy (see above). Conversion
factors between protein content and biomass dw were determined by
dividing the biomass dw by the protein content.
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RESULTS |
Batch experiments.
The degradation of propionate by a pure
culture of S. fumaroxidans in the presence of sulfate is
shown in Fig. 1A. Propionate was
stoichiometrically converted (Table 1) to
acetate while sulfate was reduced. However, only part of the propionate
was degraded. H2 accumulated to about 7 Pa and remained at
this level. A syntrophic culture of S. fumaroxidans and
M. hungatei (syntrophic coculture) degraded propionate
completely to acetate and CH4 (Fig.
2A). During the incubation,
H2 transiently accumulated to about 9 Pa but decreased at
the end to 0.8 Pa. A coculture grown on propionate plus sulfate (sulfidogenic coculture) degraded propionate almost completely to
acetate (Fig. 3A). During the incubation,
H2 accumulated to about 4 Pa but again decreased at the end
to 1.5 Pa.
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FIG. 1.
Batch monoculture of S. fumaroxidans growing
on propionate plus sulfate. (A) Changes in propionate ( ), acetate
( ), hydrogen ( ), and sulfate ( ). (B) Actual Gibbs free-energy
changes of fermentative propionate oxidation () and sulfidogenic
propionate oxidation ().
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TABLE 1.
Reactions involved in the degradation of propionate and
their standard Gibbs free-energy changes, corrected for a pH of 6.9 and a temperature of 37°Ca
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FIG. 2.
Batch syntrophic coculture of S. fumaroxidans
and M. hungatei growing on propionate. (A) Changes in
propionate (), acetate (), hydrogen (), and methane ( ). (B)
Actual Gibbs free-energy changes of fermentative propionate oxidation
(), hydrogenotrophic methanogenesis (), and syntrophic propionate
oxidation ().
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FIG. 3.
Batch sulfidogenic coculture of S. fumaroxidans and M. hungatei growing on propionate plus
sulfate. (A) Changes in propionate (), acetate (), hydrogen
(), sulfate (), and methane (). (B) Actual Gibbs free-energy
changes of fermentative propionate oxidation (),
hydrogenotrophic methanogenesis (), and sulfidogenic propionate
oxidation ().
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The actual Gibbs free energies of the degradation of propionate by a
pure culture of
S. fumaroxidans in the presence of sulfate
are depicted in Fig.
1B for both fermentative and sulfidogenic
propionate degradation. The Gibbs free-energy values of propionate
degradation under fermentative, syntrophic, and sulfidogenic
conditions,
as well as H
2-dependent methanogenesis, are
shown in Fig.
2B and
3B for the syntrophic and sulfidogenic cocultures,
respectively.
S. fumaroxidans grown in a coculture with
M. hungatei on propionate
plus sulfate is referred to as a
sulfidogenic
coculture.
The µ
max, Y
Xtot-S, and carbon and electron
recoveries were calculated for three batch cultures and are summarized
in Table
2. The culture conditions, i.e.,
pure culture, syntrophic coculture,
and sulfidogenic coculture,
significantly affected the µ
max and
Y
Xtot-S
of the cocultures. The highest µ
max value was found in
the syntrophic cocultures, followed by the sulfidogenic coculture.
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TABLE 2.
µmax, YXtot-S, C and
e recoveries, and protein conversion factora for
batch cultures of S. fumaroxidans grown in a monoculture or
in a coculture with M. hungatei on either propionate alone
or propionate plus sulfateb
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The calculated
YMPOB-S of
S. fumaroxidans and Y
Mhun-P of
M. hungatei are
presented in Table
3. The error
associated with
the calculation of the
Xi of the
propionate-degrading and methanogenic
microorganisms calculated from
the
Xtot, the N
i, and the
xi was,
in all cases, <20%. Furthermore, the
qmax values for propionate
were calculated from
µ
max and
YMPOB-S.
S. fumaroxidans obtained
the highest
YMPOB-S
in the pure-culture incubation, followed by
the syntrophic coculture
and the sulfidogenic coculture (Table
3).
M. hungatei, on
the other hand, reached the highest
YMhun-P in
the sulfidogenic coculture rather than in the syntrophic coculture
(Table
3). The highest
qmax values were observed
in the syntrophic
coculture, followed by the sulfidogenic coculture and
the pure
culture (Table
3).
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TABLE 3.
YMhun-P,
YMPOB-S, qmax, actual
G, and
YMPOB G for
propionate-oxidizing S. fumaroxidans grown under three
different batch conditions
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Chemostat experiments.
Syntrophic cocultures of S. fumaroxidans and M. hungatei were grown in chemostats
with or without a flushed headspace. The conversion of substrates to
products was generally well balanced (C balance, 116 to 130% and e
balance, 96 to 105%). The steady-state partial pressures of
H2 were 0.2 to 0.3 and 4.5 to 10 Pa in the flushed and
nonflushed systems, respectively.
The actual Gibbs free energy of propionate fermentation (reaction 1 in
Table
1) under steady-state conditions decreased linearly
(becoming
more negative) with increasing growth rate (Fig.
4A).
Due to the lower H
2
partial pressure, the Gibbs free energy was
more negative in the
flushed system than in the nonflushed system.
The ranges of Gibbs free
energies available under steady-state
conditions in the flushed and
nonflushed systems were
35.0 to
37.8 and
6.3 to
11.5 kJ per mol
of propionate, respectively.
On the other hand, the actual Gibbs free
energy of hydrogenotrophic
methanogenesis (reaction 2 in Table
1) under
steady-state conditions
increased linearly with the growth rate (Fig.
4B). Due to the
lower H
2 partial pressure, the Gibbs free
energy was more positive
in the flushed system than in the nonflushed
system. The ranges
of Gibbs free energies available under steady-state
conditions
in the flushed and nonflushed systems were
9.0 to
14.1
and
25.7
to
34.0 kJ per mol of CH
4, respectively.
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FIG. 4.
Actual Gibbs free-energy changes available for S. fumaroxidans (A) and M. hungatei (B) determined during
syntrophic growth in propionate-limited chemostat cocultures with a
flushed ( ) or nonflushed ( ) gas headspace.
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Xtot,
qs, and
qCH4 increased linearly with growth rates (data
not shown).
Xtot values were higher in the
flushed system than
in the nonflushed system. The opposite was true for
the specific
propionate conversion rates, which were the highest in the
nonflushed
system.
Growth yields of the total coculture and of each syntrophic partner
were determined individually by the total dw, the relative
cell number,
and the specific cell masses. Total dw was determined
by multiplying
the measured protein concentration by the determined
conversion factor
(Table
2).
The average growth yields (
n = 4) for the total
coculture obtained in the flushed and nonflushed systems were 4.0 ± 0.5 and
2.2 ± 0.2 g (dw) mol of S
1,
respectively. These values were used to calculate
Ytotmax and
ms-tot from the regression parameters of the
Pirt equation
(equation 4) and are listed in Table
4.
The average growth yields (
n = 4) for
S. fumaroxidans obtained in the flushed and nonflushed systems were
2.7 ± 0.5 and 1.5
± 0.1 g (dw) mol of
S
1, respectively. These values were used to calculate
YMPOBmax and
ms from the regression parameters of the Pirt
equation (Fig.
5A) and are listed in
Table
4.
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FIG. 5.
Reciprocal plots of growth yields (1/Y)
versus growth rates (1/µ) of S. fumaroxidans (A)
(1/YMPOB) and M. hungatei (B)
(1/YMhun) in propionate-limited chemostat
cocultures with a flushed () or nonflushed () gas headspace.
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The average growth yields (
n = 4) for
M. hungatei were calculated by relating the increase in the biomass
concentration to
the amount of methane formed. The amount of methane
formed was
calculated by assuming that 0.75 mol of methane was formed
per
mol of propionate (Table
1, reaction 3). The growth yields of
M. hungatei obtained in the flushed and nonflushed systems
were
6.6 ± 0.6 and 4.6 ± 0.7 g (dw) mol of
P
1, respectively. These values were used to calculate
YMhunmax and
ms from the regression parameters of the Pirt
equation (Fig.
5B) and are listed in Table
4.
Ds01/
rAx,
calculated from the determined
Ytotmax of the chemostat cocultures
of
S. fumaroxidans and
M. hungatei, in the
flushed
and nonflushed systems, were 56.1 and 167.6 kJ mol of
C
1, respectively. The
Ds01/
rAx
calculated from the determined
YMPOBmax of
S. fumaroxidans in the flushed and nonflushed systems were
320.7
and
705.7 kJ mol of C
1,
respectively.
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DISCUSSION |
Energetics of syntrophic propionate oxidation.
The Gibbs free
energy available for propionate-oxidizing S. fumaroxidans
was found to be higher in the flushed (35 to 38 kJ mol of
propionate1) than in the nonflushed (6 to 12 kJ mol
of propionate1) chemostat cocultures. A possible
disequilibrium of H2 between the liquid and gas phases
would have resulted in even higher Gibbs free energies (less exergonic)
than indicated by the values above. Disregarding this potential bias,
the apparent Gibbs free-energy change in the flushed system is
sufficient to generate not more than 1/2 mol of ATP, while in the
nonflushed system less than 1/11 to 1/7 mol of ATP can be generated.
The possible ATP generation based on the Gibbs free-energy change in
the syntrophic batch cultures (12.6 kJ mol of
propionate1; Table 3) was similarly low as in the
nonflushed chemostat. The Gibbs free-energy change in the flushed
chemostat system is more than the minimum energy quantum necessary to
sustain microbial life (23 kJ mole of substrate1 
1/3 mol of ATP), but this is not the case in the nonflushed chemostat
and in the batch culture. Nevertheless, our experiments show that
microbial growth was sustained even in the nonflushed system. Hence,
the available Gibbs free energy was apparently sufficient. Relatively
small free-energy changes during the degradation of propionate have
also been reported for different methanogenic environments (5, 19,
26, 46). These observations raise the question of how S. fumaroxidans and other syntrophic propionate-oxidizing bacteria
manage to exploit the little Gibbs free energy available for the
generation of ATP. More research is required to answer this question.
Growth parameters.
Measured values of growth parameters
obtained in batch experiments depended on the type of culture and the
overall propionate-consuming reaction. S. fumaroxidans had
the highest YMPOB-S but the lowest µmax when grown as a pure culture (propionate plus
sulfate). The values of YMPOB-S and
µmax (Table 2) we obtained correspond well to the
previously reported values of 1.5 g (dw) mol of S1
and 0.024 day1 (40), respectively. The
µmax values reported for S. fumaroxidans cocultured with M. hungatei on propionate was reported to be
0.17 day1 (32). We obtained a value of
0.21 ± 0.01 day1 for µmax, which is
in the same range as the reported value. Growth yields of S. fumaroxidans in syntrophic or sulfidogenic cocultures are not
reported in the literature. The growth yields we calculated for
S. fumaroxidans were 1.02 ± 0.04 and 0.82 ± 0.04 g (dw) mol of S1, respectively. Our
results showed that the calculated and reported yields of S. fumaroxidans grown as a pure culture were higher than the values
obtained in syntrophic and sulfidogenic cocultures. The reason for the
lower growth yields in the cocultures can be explained by the fact
that the available energy has to be shared between the two syntrophic partners.
The
YMPOBmax and the
ms for
S. fumaroxidans and
M. hungatei cocultured on propionate were determined in chemostat
cultures (Table
4). To our knowledge, these are the first data obtained
for a
syntrophic propionate-oxidizing bacterium. The values are
comparable
to those determined for
Pelobacter acetylenicus
growing on ethanol
syntrophically with different
H
2-consuming anaerobes (
31).
Maintenance energy.
Tijhuis et al. (38) showed that
the maintenance requirements of microorganisms can be described on the
basis of mE, which theoretically should only be
a factor of temperature. Thus, the theoretical
mE values at 28, 30, and 37°C are 4.4 ± 1.4, 5.2 ± 1.7 and 9.8 ± 3.1 kJ mol of biomass
C1 h1, respectively (38). We
calculated the parameter mE from literature data
on an ethanol-oxidizing syntrophic coculture on several anaerobic pure
cultures and included the data obtained from the chemostat cocultures
of S. fumaroxidans and M. hungatei (Table
5). Interestingly, these
mE values were usually lower than the
theoretical values, some by more than 1 order of magnitude.
View this table:
[in this window]
[in a new window]
|
TABLE 5.
mE determined for different
anaerobic monocultures and cocultures grown under different conditions
using the measured ms and the
Gav01 of the overall
catabolic reaction
|
|
Heijnen and VanDijken (
13) pointed out that to obtain
meaningful values, the
Gav should be
calculated from the actual concentrations
of reactants and products and
that the use of
Gav01, as done
for calculation of the data in Table
5, is only a compromise
if actual
concentrations are not available. Therefore, we also
calculated
mE values from the actual
Gav values measured for
propionate oxidation
by
S. fumaroxidans (Fig.
4A) and
H
2-CO
2-dependent
methanogenesis by
M. hungatei (Fig.
4B) in the chemostat experiments,
using the
species-specific
ms values (Table
6). Again, however,
the
mE values of
S. fumaroxidans and
M. hungatei were more than
1 order of magnitude lower than
the theoretical values expected
from the equation of Tijhuis et
al. (
38). Obviously, the maintenance
energies
calculated from our chemostat experiments and from earlier
experiments
with anaerobic cultures are not consistent with the
pertinent theory,
suggesting that microorganisms, at least anaerobes,
are able to grow at
maintenance energies lower than those theoretically
predicted. Tijhuis
et al. (
38) have concluded that (i) the electron
acceptor,
(ii) different C sources, (iii) the type of organism,
and (iv) the use
of mixed sludges versus pure cultures have no
effect on
mE. However, this conclusion was based on
limited culture
data. In particular, syntrophic cultures have not been
included.
In addition, Heijnen and VanDijken (
13)
cautioned that the derived
theoretical relationships give only a first
approximation based
on thermodynamics and that modifications may be
necessary if mechanistic
details become available, e.g., microbes
exploiting the available
Gibbs free energy by using different metabolic
pathways with different
energetic efficiencies. There are even
differences among the various
physiological groups of anaerobic
microbes in whether they depend
on changes in reaction enthalpy or
reaction entropy (
42). For
example,
H
2-CO
2-dependent methanogenesis depends mainly
on the
enthalpy change and is retarded by the entropy change whereas
anaerobes, syntrophs in particular, depend mainly on the entropy
change
(
42). Apparently, more research is necessary to elucidate
the theoretical background of maintenance energy in fastidious
anaerobic bacteria.
View this table:
[in this window]
[in a new window]
|
TABLE 6.
mE determined from the
ms and the actual Gav measured
for S. fumaroxidans and M. hungatei grown in
syntrophic cocultures with two different chemostat setups
|
|
Gibbs free-energy dissipation per mol of biomass C.
Heijnen
and VanDijken (13) showed that
Ds01/rAx can
be regarded as a simple thermodynamic measure of the amount of
biochemical "work" required to convert a carbon source into biomass
and can be used to characterize chemotrophic microbial growth. We
calculated Ds01/rAx from
the maximum growth yields determined for the syntrophic chemostat
cocultures of S. fumaroxidans and M. hungatei.
The calculated Ds01/rAx
values for the flushed and nonflushed systems were 56.1 and 167.6 kJ
mol of biomass C1, respectively. These values are at the
lower end of the range of values (150 to 3,500 kJ mol of biomass
C1) observed for various modes of chemotrophic growth
(13). Furthermore, we calculated
Ds01/rAx for
S. fumaroxidans alone and obtained values of 320.7 and 705.7 kJ mol of biomass C1, respectively, for the
flushed and nonflushed systems. These negative values are not realistic
and can be explained by the fact that the
Ds01 is endergonic when using the
standard Gibbs energies of formation for the reactants and products
under standard conditions.
Therefore, we also calculated theoretical growth yields from a
theoretical
Ds01/
rAx,
which is 426 kJ mol of biomass C
1 for propionate. For the
syntrophic coculture (equation 7), we
calculated a
Ytotmax of 4.4 g mol of
S
1, i.e., similar to the actually observed values (Table
4). However,
for
S. fumaroxidans alone (equation 8), we
obtained a negative
YMPOBmax value,
indicating that the theoretical
Ds01/
rAx must
be too low. Only when we assumed a
Ds01/
rAx of
>622 kJ mol of biomass C
1 did the calculated
YMPOBmax become positive. When
assuming a
Ds01/
rAx of
3,500 kJ mol of biomass C
1, we calculated a
YMPOBmax of 1.95 g mol of
S
1, i.e., similar to the actually observed values (Table
4). A
Ds01/
rAx as
high as 3,500 kJ mol of biomass C
1 is usually observed in
chemolithoautotrophic bacteria that use
CO
2 as a carbon
source and require the occurrence of reversed
electron transport, e.g.,
nitrifiers and thiobacilli. Our data
indicate that syntrophic
propionate oxidation by
S. fumaroxidans falls into the same
category of Gibbs free-energy
dissipation.
Energetic efficiency of growth.
We were able to determine both
the growth yield of S. fumaroxidans and the Gibbs free
energy available by oxidation of propionate (Table 3). Although the net
generation of ATP during the degradation of propionate is not clear, we
were able to use the determined yields and Gibbs free-energy values to
estimate a proxy (i.e., YMPOBG) for
YATP, i.e., the biomass synthesized from 1 mol
of ATP generated. Under physiological conditions, an average Gibbs free
energy of 70 kJ is needed for the irreversible synthesis of 1 mol of
ATP. Therefore, YMPOBG
was calculated by normalizing the measured
YMPOB-S to an energy quantum of 70 kJ
mol1 (16, 31). The results are shown in Table
3.
The
YMPOBG values
calculated for
S. fumaroxidans grown on propionate plus
sulfate in either a monoculture and a sulfidogenic
coculture were 1.2 and 0.9 g (dw) 70 kJ
1, respectively. These values
are much lower than the values of
5 to 12 g mol of
ATP
1 reported by Stouthamer (
34) for
fermenting bacteria. The
YMPOBG value for
S. fumaroxidans grown on propionate in a syntrophic
coculture was 5.7 g (dw) 70 kJ
1. The
YMPOBG values
calculated for flushed and nonflushed chemostat cocultures
grown on
propionate were 4.5 to 6.2 and 10.3 to 15.7 g (dw) 70
kJ
1, respectively. These values suggest that the energy
efficiency
in syntrophic cocultures was much higher than in
sulfidogenic
monocultures and
cocultures.
A possible reason for the energetic inefficiency of sulfidogenic
oxidation of propionate may be found in the mechanism of
the reduction
of sulfate to sulfide. With sulfate as an electron
acceptor, substrate
level phosphorylation might be more effective
for the syntrophic
propionate oxidizer but sulfate has to be activated
first. This
activation costs the cell 2 mol of ATP per mol of
sulfate.
Additionally, the cell losses another 1/3 mol of ATP
during the
transport of sulfate across the microbial membrane.
So, the activation
and transport of sulfate costs the cell 2 1/3
mol of ATP per mol of
sulfate (
47). This amount of energy cannot
be generated at
substrate level phosphorylation alone (maximum
of 1 1/3 mol of ATP per
mol of sulfate consumed). Thus, it is
most likely that
S. fumaroxidans associates the reduction of sulfate
with an electron
transport chain allowing chemiosmotic ATP synthesis.
If this process
operates inefficiently, it may explain the observed
low energy
efficiency for growth. Furthermore, this hypothesis
could explain why
syntrophic propionate-oxidizing bacteria are
outcompeted by
propionate-oxidizing sulfate reducers. However,
to our knowledge, no
data concerning the energy efficiency for
growth on propionate and
sulfate for propionate-oxidizing sulfate
reducers is available. Thus,
there is no direct evidence to support
these
hypotheses.
Conclusion.
It is postulated that the small Gibbs free-energy
changes observed during the degradation of propionate in different
methanogenic environments are sufficient to sustain microbial growth.
However, the Gibbs free-energy changes observed are much lower than
those theoretically predicted. Furthermore, it is assumed that the
calculated low energetic efficiency of S. fumaroxidans
during growth on propionate plus sulfate can be due to an inefficient
electron transport chain involved in chemiosmotic energy conservation.
The observed growth yields further suggest that S. fumaroxidans requires a relatively large Gibbs free-energy
dissipation for biomass synthesis, similar to that typically observed
for chemolithoautotrophic bacteria with reversed electron transport.
| |
ACKNOWLEDGMENT |
This work was financially supported by the Fonds der Chemischen
Industrie, Germany.
| |
FOOTNOTES |
*
Corresponding author. Mailing address:
Max-Planck-Institut für terrestrische Mikrobiologie,
Karl-von-Frisch-Str., D-35043 Marburg, Germany. Phone: 49 (6421) 178 801. Fax: 49 (6421) 178 809. E-mail:
conrad{at}mailer.uni-marburg.de.
Present address: Department of Biological Sciences, University of
Warwick, Coventry CV4 7AL, United Kingdom.
| |
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Abstract
Introduction
