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Keywords:

  • Competition;
  • Denitrification;
  • Inhibition;
  • Methanogenesis;
  • Most-probable-number enumeration;
  • Sulfate reduction

Abstract

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Methods
  5. 3Results
  6. 4Discussion
  7. Acknowledgements
  8. References

Acetate is quantitatively the most important substrate for methane production in a freshwater sediment in The Netherlands. In the presence of alternative electron acceptors the conversion of acetate by methanogens was strongly inhibited. By modelling the results, obtained in experiments with and without 13C-labelled acetate, we could show that the competition for acetate between methanogens and sulfate reducers is the main cause of inhibition of methanogenesis in the sediment. Although nitrate led to a complete inhibition of methanogenesis, acetate-utilising nitrate-reducing bacteria hardly competed with methanogens for the available acetate in the presence of nitrate. Most-probable-number enumerations showed that methanogens (2×108 cells cm−3 sediment) and sulfate reducers (2×108 cells cm−3 sediment) were the dominant acetate-utilising organisms in the sediment, while numbers of acetate-utilising nitrate reducers were very low (5×105 cells cm−3 sediment). However, high numbers of sulfide-oxidising nitrate reducers were detected. Denitrification might result in the formation of toxic products. We speculate that the accumulation of low concentrations of NO (<0.2 mM) may result in an inhibition of methanogenesis.


1Introduction

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Methods
  5. 3Results
  6. 4Discussion
  7. Acknowledgements
  8. References

Acetate is quantitatively the most important substrate for methanogens in methanogenic environments [1–4]. Up to 70–80% of the methane in freshwater sediments may be derived from acetate; the remainder is formed by reduction of bicarbonate with hydrogen. The consumption of acetate or hydrogen by methanogens is strongly affected by the competition with anaerobic respiring microorganisms. Consequently, one of the factors regulating methane formation in nature is the availability of inorganic electron acceptors such as nitrate, sulfate, sulfur and oxidised metals (FeIII, MnIV) [2].

Insight into the effect of sulfate on H2-dependent methanogenesis has been obtained in sediment studies and in studies with pure cultures of H2-consuming methanogens and sulfate reducers [5–8]. It was shown in batch cultures that sulfate reducers outcompete methanogens for H2 due to a higher affinity and higher growth yield [5,8]. Unfortunately, less is known about the competition between methanogens and sulfate reducers for acetate. Schönheit et al. [9] showed that Desulfobacter postgatei has a higher affinity for acetate than Methanosarcina barkeri. This could explain why Desulfobacter species are the main acetate-degrading microorganisms in marine sediments. However, Methanosaeta rather than Methanosarcina species are the dominant acetate-degrading methanogens in various methanogenic environments [3,10]. These methanogens display maximum acetate uptake rates (Vmax), half-saturation constants (Km) and maximum growth rates (μmax) similar to acetate-utilising sulfate reducers isolated from freshwater environments [11,12]. Thus, the effect of sulfate on the fate of acetate is not as clear-cut as its effect on the fate of H2.

Nitrate is also known to suppress methane formation [13–16]. It was speculated that production of the toxic intermediates of denitrification (nitrite, NO, N2O), rather than the competition for acetate between methanogens and denitrifiers, was responsible for the inhibition of methanogenesis [13,17–19]. Remarkably, little is known about the role of acetate as electron donor for nitrate reduction in natural environments. Most nitrate-reducing bacteria have only been tested for their capacity to grow on acetate and nitrate. Information about their growth kinetic properties on acetate is scarce. Quantification and identification of acetate-utilising denitrifiers may give insight into their role in acetate metabolism in anoxic environments.

We analysed the effect of sulfate and nitrate on methane formation in freshwater sediment by using 2-13C-labelled acetate. To obtain insight into the effect of inorganic electron acceptors on potential methanogenesis, we quantified the acetate-utilising methanogens, sulfate reducers and nitrate reducers in the sediment. Furthermore, a mathematical model was used to analyse possible interactions between the different microbial populations in the sediment.

2Methods

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Methods
  5. 3Results
  6. 4Discussion
  7. Acknowledgements
  8. References

2.1Site description and sample collection

The polder Zegvelderbroek was chosen as the study area and the polder is representative for other polders in The Netherlands. In previous centuries the peat lands in this area were largely drained and reclaimed, and divided into separate polders. Peat extraction for fuel was practised at a large scale [20,21]. In this century water management of the polders was profoundly intensified in favour of agriculture. Today, it is a peat grassland area in which ditches are lying at roughly every 40–100 m. Between the ditches lie small long strips of grassland and smaller cross-ditches [22].

Sediment samples were collected on 4 September 1995 (summer) and 4 April 1996 (spring) from ditches in the polder Zegvelderbroek [23]. The samples were collected with a sediment corer as described previously [23]. The temperature of the sediment in September 1995 and April 1996 was 13 and 7°C, and that of the water 14 and 6°C, respectively. The cores were stored at 10°C. Four days after sampling the sediments were processed further. The upper 0–10 cm of the sediment were used in the experiments. This part of the sediment had a water content of 88–95% by weight, and a density of 1.12–1.15 g cm−3. Dry sediment material (peat) had an organic content of 90–95% (n=5).

Pore water samples were taken from the top 12 cm of the core on the day of sampling. The samples were drawn by 1-ml syringe through 6-mm holes (covered with Scotch tape No. 471) in the core at different depth intervals. For analyses of anions, the samples were centrifuged for 15 min at 17 380×g. Supernatants were stored at −20°C and analysed by high performance liquid chromatography (HPLC).

2.2Media

A basal bicarbonate-buffered medium described by Huser et al. [24] was used for most-probable-number (MPN) counts. To 1 l of medium, 1 ml of a vitamin solution [25] and 1 ml each of an acid and an alkaline trace elements solution [26] were added. The vitamin solution was filter-sterilised separately. The gas phase above the medium was 172 kPa N2/CO2 (80%/20%) resulting in a medium pH of 6.8–6.9. Acetate, sulfate and nitrate were added from 1-M heat-sterilised stock solutions.

2.3Quantification of functional groups of bacteria

All manipulations were done under anaerobic conditions in a glove box. The 0–10-cm layer of the sediment was homogenised, and 15 ml were transferred to a 250-ml serum bottle containing 135 ml of medium. The bottle was closed with a butyl rubber stopper, evacuated and gassed with N2/CO2 (80%/20%). After shaking the bottle for 5 min, the sediment slurry (15 ml) was serially diluted to a 10−10 dilution. A three-bottle MPN series was prepared by transferring 5-ml samples to 120-ml serum bottles containing 45 ml of medium. The MPN tests for acetate-utilising bacteria were performed with medium containing 10-mM acetate with or without sulfate or nitrate (10 mM). Incubations were carried out in the dark at 20°C. Bottles were examined weekly and were considered negative if no growth occurred after 6–12 months of incubation. The concentrations of the substrate and products were determined in positive bottles. Transfer to fresh medium confirmed growth in the highest positive dilution. MPN, standard deviation and 95% confidence intervals were determined using the computer program of Hurley and Roscoe [27]. Enumerations in the summer and spring samples were evaluated with Student's t-test (α<0.05) using exponentially transformed bacterial counts. The populations were expressed as cells per cm3 of wet sediment. An aceticlastic methanogen and sulfate reducer were previously isolated from the MPN series [28].

2.4Acetate consumption in resting cell suspensions

The Michaelis–Menten kinetic parameters Vmax and Km for the isolated aceticlastic methanogen and sulfate reducer were estimated from acetate depletion curves (acetate consumption versus time) obtained with concentrated cell suspensions as described by Oude Elferink et al. [12]. The depletion data were fitted to a linearisation of the Michaelis–Menten model [29]. The specific cell mass (bi: g dry weight (dw) cell−1) of the isolated aceticlastic methanogen and sulfate reducer was determined as described by Seitz et al. [30].

2.5Setup of incubation experiments

An optimal experimental design of the tracer experiments was determined using a statistical program (CADEMO) [31]. Every experiment was performed with a total set of n=18 (incubations without acetate addition) or n=21 (incubations with acetate addition) tubes. At each sampling point in time a number of tubes (r) were sacrificed for gas (r=3 for all incubations) or liquid analysis (r=3 for incubations without acetate addition and r=4 for acetate incubations).

All manipulations were done under anaerobic conditions in a glove box. Homogenised sediment from the 0–10-cm layer was distributed in 10-ml portions into 26-ml tubes and closed with butyl rubber stoppers. The tubes were seven times evacuated and gassed with N2 (152 kPa), and stored at 10°C. The next day [2-13C]acetate and electron acceptors were added. Labelled acetate was added to a final concentration of 120–150 μM. Controls without added acetate were prepared as well. Inactivated controls were made by adding formaldehyde to a concentration of 3.75%. The total recovery of labelled acetate from these tubes was higher than 90%. All tubes were incubated in the dark at 17°C. Tubes were acidified with 3 M HCl to stop microbial activity, and then gas samples were taken by syringe from the headspaces and analysed by gas chromatography (GC) for CH4 and CO2. For analysis of [2-13C]acetate, anions and other dissolved intermediates, the contents of the tubes were centrifuged two times for 15 min at 17 380×g. Supernatants were stored at −20°C and analysed by GC and HPLC. The pH was checked at the end of the experiment and was always between 6.8 and 7.0.

2.6Calculation of rate constants and turnover rates

Rate constants of consumption of 13C-acetate, sulfate and nitrate or production of 13C-CH4 or sulfate were computed by regression of the natural logarithm of concentration versus time as described by De Graaf et al. [32]. It is assumed that the microbial biomass is constant during the incubation. The 13C-labelled carbon is converted into CH4 or CO2 but not incorporated in the biomass. The rate constant of methanogenic acetate consumption was calculated by assuming that the acetate consumed by the methanogens may be described as:

  • image(1)

where [13C-AcMPB]t=x is the amount of 13C-acetate consumed by the aceticlastic methanogens. The rate constant of the sulfidogenic acetate consumption was calculated by assuming that the acetate consumed by the sulfate reducers may be described as:

  • image(2)

where [13C-AcSRB]t=x is the amount of 13C-acetate consumed by the aceticlastic sulfate reducers and [13C-AcMPB]tot the total amount of 13C-acetate consumed by the aceticlastic methanogens at the end of the incubation (see Eq. 1).

Rates of consumption and production (V: mol l−1 h−1) were calculated by multiplying rate constants of consumption for 13C-acetate or electron acceptors, or production of 13C-CH4 or sulfate with the actual pool sizes at time zero. For the calculation of the 13C-CH4 production rate we used the actual pool size of total CH4 (12C/13C) at time zero. To compare the rates of consumption or production we performed Student's t-tests (α<0.05).

2.7Calculation of cell numbers based on consumption rates

The numbers of acetate-utilising sulfate-reducing and methanogenic cells were calculated from their actual activities. In kinetic studies where bacterial growth can be neglected, Michaelis–Menten kinetics may be used to describe the substrate uptake rate of whole cells:

  • image(3)

where Vxmax and Vx are given in moles per gram of cellular dry weight per hour (mol (g dw)−1 h−1). The biomass specific activity of the sulfate reducers or methanogens can be obtained from the consumption rates (V) of acetate (see Section 2.6) divided by the total cell mass of each population, Bi ((g dw) l−1):

  • image(4)

The dry mass of a microorganism may be described as:

  • image(5)

where Ni is the number of cells in the sediment (cells l−1) and bi is the specific cell mass ((g dw) cell−1). By determining the Vmax and Km values, and specific cell masses of the dominant acetate-consuming sulfate reducers and methanogens (see Section 2.4) the number of cells based on the acetate consumption rates can be calculated according to:

  • image(6)

These values based upon activity measurements could be compared with the numbers obtained by MPN counts.

2.8Model structure

A model was developed to describe and explain the influence of alternate electron acceptors on methane production. A full description and validation of the model were presented previously [33]. The alternative electron acceptors incorporated in the model are nitrate and sulfate. In the model, acetate is considered to be the only carbon substrate. All reactions are described by Michaelis–Menten kinetics. In this way, the competition can be described from the maximum rate (Vmax) and the affinity constants (Km) for electron acceptors and electron donors. In principle, Vmax is a function of the microbial biomass. However, in our model, it is assumed that the microbial biomass is constant in time [34,35]. In the short-term incubations described here (6–7 h), little growth of microbial biomass will occur. Affinity constants and potential reaction rates were derived from data obtained in this study and from the literature (Table 1). In addition, non-competitive inhibition factors for the inhibition of methane production by sulfide, NO and N2O, inhibition of sulfate reduction by sulfide and the inhibition of nitrate reduction by sulfide were incorporated and parameterised using literature data. With the description of the freshwater sediment thus obtained, the influence of different reactions was determined with a sensitivity analysis. With such an analysis a better understanding of the processes in the sediment can be obtained.

Table 1.  Parameters derived from this study and the literature, which were used for the modelling
  1. aDerived from acetate consumption in resting cell suspensions of the most dominant acetate-utilising methanogen and sulfate reducer isolated from the sediment.

  2. bDerived from sediment incubations by parameterisation of the model.

  3. cTaken from van Bodegom and Scholten [33].

ReactionKm electron donor (mM)Km electron acceptor (mM)Vmax (mM s−1)
Acetate+H2O[RIGHTWARDS ARROW]CH4+HCO30.46a9.5×10−4b
Acetate+SO42−[RIGHTWARDS ARROW]2 HCO3+HS0.51a0.23c5.7×10−4b
5/4 Acetate+2 NO3+3/4 H+[RIGHTWARDS ARROW]21/2 HCO3+N2+H2O0.09c0.42c1.2×10−4c

2.9Analytical techniques

Determination of 13C-labelled acetate, methane and carbon dioxide was carried out on a GC (Hewlett Packard model 5890) equipped with a mass-selective detector (HP model 5971A). Acetate and its stable isotopes were analysed with a capillary column (Innowax, Hewlett Packard, Amsterdam, The Netherlands) and their detection limit was 5 μM. Methane, carbon dioxide and their stable isotopes were separated on a capillary plot-fused silica column (coating Poraplot Q, Chrompack, Middelburg, The Netherlands). Total methane was measured on a 417 Packard chromatograph equipped with a flame ionisation detector (FID) and a molecular sieve 5A column. Anions (chloride, nitrate, nitrite, sulfate and thiosulfate) were analysed by HPLC as described previously [23]. Sulfide was determined as described by Trüper and Schlegel [36]. Samples for the analysis of sulfide were kept on ice in closed Eppendorf tubes and determined at the end of the experiment. Determination of methane and volatile fatty acids in the MPN incubations was done as described previously [28]. Organic acids (including acetate) in the sediment incubations and profiles were determined as described by Scholten et al. [4]. Hydrogen was determined as described by Houwen et al. [37].

3Results

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Methods
  5. 3Results
  6. 4Discussion
  7. Acknowledgements
  8. References

3.1Pore water profile

Variations in sulfate concentrations in the pore water were related with depth and season (Fig. 1). In summer and spring, sulfate concentration gradients showed strong decreases in the first 0.5 cm of the sediment. Because vertical convection was negligible, the steep concentration gradients of sulfate indicated the occurrence of sulfate reduction. Below 5 cm the sulfate concentrations increased again. Nitrate and nitrite concentrations were below the detection limit (<1 μM) in the summer and spring profiles. Organic acid concentrations in the summer and spring gradients were below the detection limit (<10 μM).

image

Figure 1. Sulfate concentration profiles at the sediment surface during summer (September 1995, ▪) and spring (April 1996, ▴).

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3.2Sediment incubations

Effects of sulfate and nitrate on the biotransformation of 13C-labelled acetate were studied. During the 6 h of incubation, almost no methane was produced in the sediment without acetate added (Fig. 2A–C). Addition of [2-13C]acetate to sediment collected in summer stimulated the production of methane (Fig. 3A). Aceticlastic methanogenesis accounted for more than 70% of the conversion of labelled acetate (Fig. 3A). Sulfate reduction was not stimulated by the addition of labelled acetate alone, although 50–70 μM sulfate was present in the pore water (Fig. 3A). However, addition of labelled acetate in combination with sulfate inhibited the production of labelled methane (Fig. 3B and Table 2). The consumption rate of sulfate was increased by the addition of acetate (Table 2). The sulfate-reducing community was limited by the availability of sulfate in the sediment incubations (Fig. 3B). Addition of nitrate, with or without labelled acetate, led to the accumulation of sulfate (Figs. 2C and 3C). Methanogenesis was inhibited by the activity of nitrate-reducing bacteria but because sulfate accumulated it remained unclear whether the sulfate-reducing activity was inhibited as well. The model underestimated the depletion of nitrate in the incubations without labelled acetate (Fig. 2C) and overestimated the depletion of nitrate in the incubations with labelled acetate (Fig. 3C). Remarkably, the acetate consumption rate decreased by the addition of nitrate (Table 2). However, acetate addition did not affect the consumption rate of nitrate, suggesting that nitrate-reducing bacteria showed no additional response to acetate. The model overestimated the depletion of acetate in the incubations with labelled acetate and nitrate (Fig. 3C). The accumulation of sulfate seems to indicate that reduced sulfur compounds served as electron donors for the denitrifying population. The model was not able to describe the accumulation of sulfate in the incubations with nitrate (Figs. 2C and 3C).

image

Figure 2. The effect of sulfate or nitrate on CH4 formation in anaerobic freshwater sediment. A: Non-supplemented sediment; B: sediment supplemented with sulfate; C: sediment supplemented with nitrate. Measured (open symbols) and modelled (lines) concentrations: ◯/dot, sulfate; □/solid, total CH4; ⋄/short dash, nitrate. Error bars represent 1 S.E.M. (r=3) for gas and liquid analysis. For an explanation of r see Section 2.

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image

Figure 3. The effect of sulfate or nitrate on CH4 formation in anaerobic freshwater sediment. A: Supplemented with 13C-acetate; B: supplemented with 13C-acetate and sulfate; C: supplemented with 13C-acetate and nitrate. Measured (open symbols) and modelled (closed symbols+lines) concentrations: ▵/dash dot dot, acetate; ◯/dot, sulfate; □/solid, 13C-CH4; ⋄/short dash, nitrate. Error bars represent 1 S.E.M. for (r=3) gas analysis and for (r=4) liquid analysis. For an explanation of r see Section 2.

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Table 2.  Potential consumption rate for 13C-acetate, sulfate and nitrate and production rate for 13C-methane and sulfate (μmol l−1 h−1) obtained from incubations of freshwater sediment samples for 6 h (September 1995) or 7 h (April 1996) at 17°C
  1. a–pDifferent letters in a column indicate significant differences (P<0.05) in consumption or production rate constants based on Student's t-test. The standard errors (1 S.E.M.) of the consumption/production rates are shown in parentheses.

IncubationConsumption rateProduction rate
 13C-acetateSO42−NO313C-CH4SO42−
September sample
Acetate62 (10)a3 (2)e77 (21)m
Unsupplemented control3 (3)e
Acetate+SO42−61 (18)a16 (3)f24 (13)n
Acetate omitted control10 (4)g
Acetate+NO335 (11)b52 (13)k05 (2)p
Acetate omitted control61 (14)k6 (3)p
April sample
Acetate+SO42−27 (3)c26 (9)h5 (3)o
Acetate omitted control1 (9)i
Acetate+NO312 (1)d14 (7)j57 (9)l0
Acetate omitted control8 (12)j62 (11)l

The natural pool of sulfate was between 0.5 and 1.2 mM in sediment collected in spring, which is much higher than the sulfate concentration found in summer (Fig. 1). Almost no sulfate was reduced or methane produced in the sediment without substrate addition, indicating that methanogens and sulfate reducers were both limited by substrate availability (Table 2). Addition of labelled acetate instantaneously stimulated sulfate reduction. In this case, sulfate reduction accounted for more than 75% of the acetate conversion. The initial production of labelled methane was low but increased after 4 h of incubation (data not shown). Nevertheless, the production of labelled CH4 showed that aceticlastic methanogens competed with the sulfate reducers for the available acetate (Table 2). The consumption rate of acetate was again lowered by the addition of nitrate (Table 2). Determination of the contribution of sulfate reduction and nitrate reduction to the consumption of labelled acetate (13CO2 formed/SO42− reduced and 13CO2 formed/NO3 reduced) was not possible in all incubations, due to the high background concentration of 13CO2 (>50 μM). This background was caused by the acidification of our samples (see Section 2.5) which released high amounts of CO2 (>5 mM). In all summer and spring incubations the sulfide concentrations were between 0.3 and 0.5 and 0.2 and 0.4 mM, respectively, and the H2 partial pressure was below the detection limit (<20 Pa).

3.3Quantification of methanogenic, sulfate-reducing and denitrifying microorganisms

Populations of three acetate-using bacterial groups were enumerated with sediments sampled in summer and spring (Table 3). Large differences in the summer and spring samples were not observed. The incubation times required to enumerate the three types of microorganisms were different. In MPN dilutions used to enumerate acetate-utilising nitrate-reducing bacteria, the maximum cell number was already reached after 4–6 weeks, whereas in the dilution series used to enumerate acetate-utilising sulfate-reducing bacteria and acetate-consuming methanogens final cell numbers were obtained after 9–10 months. The determined values for Vmax, Km and bi of the isolated aceticlastic methanogen were: Vmax: 3.0±1.2 μmol h−1 (g dw)−1, Km: 0.46±0.16 mM, and bi: 1.0 pg. For the sulfate reducer these values were: Vmax: 1.8±0.7 μmol h−1 (g dw)−1, Km: 0.51±0.25 mM, and bi: 0.85 pg. The determination of the Vmax and Km values for the isolated acetate-utilising nitrate reducer in concentrated cell suspensions was not successful. The estimation of the number of acetate-utilising methanogens and sulfate reducers based on consumption rates of acetate are given in Table 3. The calculated numbers of acetate-utilising methanogens and sulfate reducers were in general highly similar to the numbers obtained with the MPN counts.

Table 3.  Results of MPN experiments, and estimated number of cells based on consumption rates of 13C-labelled acetate in Zegveld sediment sampled in summer (September 1995) and spring (April 1996)
  1. Experiments were performed at 17°C.

  2. a95% confidence interval in parentheses.

  3. bMinimum and maximum values in parentheses.

  4. cNT, not tested.

Acetate-utilising microorganismsCells cm−3 sediment
 MPN countsaEstimates from consumption rateb
 September sampleApril sampleSeptember sampleApril sample
Methanogens2×108 (1–8)2×108 (1–8)8×107 (5–13)4×107 (2–5)
Sulfate reducers2×108 (1–8)9×107 (3–42)2×108 (1–3)8×107 (4–12)
Nitrate reducers5×105 (2–28)9×104 (3–43)NTcNT

4Discussion

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Methods
  5. 3Results
  6. 4Discussion
  7. Acknowledgements
  8. References

The pore water profiles showed that sulfate concentrations varied with depth and season (Fig. 1). The observed steep concentration gradients of sulfate indicated that sulfate reduction occurred in the first 0.5 cm of the sediment. Below 5 cm the sulfate concentrations increased again to constant values in the bottom centimetres. This suggests that either sulfate reduction did not occur or that sulfate reduction was in equilibrium with sulfate production by chemolithotrophic denitrifiers (discussed below). Unfortunately, the measurement of sulfate reduction in situ (by tracer method) to confirm that the steep concentration gradients were indeed a result of ongoing sulfate reduction was technically not possible in this study. However, pore water profiles were made every 2 months (3–5 cores per sample date) over a 2-year period and these steep concentration gradients of sulfate were observed especially during the spring, summer and autumn [38]. During the winter months the profiles showed practically continuous concentration gradients. Remarkably, most of the sulfate reduction seems to take place in the first centimetre of the sediment. We were not able to measure the penetration depth of oxygen in the sediment but it was shown to be at a maximum of 2–3 mm in similar freshwater sediment [39]. This suggests that oxygen levels at the sediment/water interface were low enough to allow sulfate reduction to take place. In the summer and spring profiles organic acid concentrations were generally below detection limit (<10 μM) but sporadically in autumn (1994 and 1995) in the first centimetre of the sediment, formate (10 μM), acetate (300 μM) and lactate (25 μM) accumulated. This accumulation of acetate in the sediment was probably due to an increased input of dead organic matter. It justifies the addition of relatively high concentrations of 13C-acetate (120–150 μM) in our sediment incubations.

Our experiments showed that in summer, aceticlastic methanogenesis is the dominant acetate-consuming process in the freshwater sediment from Zegvelderbroek. The quantitative importance of acetate to methanogenesis could not be judged in these experiments, but an inhibition study revealed that about 70–80% of the total carbon flow to CH4 was through acetate [4]. In our experiments, methanogenesis seemed to account for more than 70% of the acetate mineralisation in the sediment when sulfate concentrations were between 50 and 70 μM. The reduction of sulfate at these low concentrations was not stimulated by the presence of acetate. However, addition of sulfate stimulated sulfate reduction, suggesting that the acetate-utilising sulfate reducers are sulfate limited at less than 70 μM. Indeed, sulfate reduction dominated the consumption of acetate (>75%) when sulfate concentrations in the sediment were higher than 500 μM (Fig. 3A,B). Stimulation of both processes by the addition of acetate showed that acetate-utilising methanogens and sulfate reducers were competing directly for the available acetate. This was confirmed by our modelling results (Fig. 3A,B). Previous studies had shown that methanogenesis and sulfate reduction were mainly confined to the first 10 cm of the sediment (Scholten, unpublished). The pore water profiles showed steep concentration gradients of sulfate, indicating that sulfate reduction occurred in the first 0.5 cm of the sediment (Fig. 1). As the sulfate concentration increased again below 10 cm it seemed that sulfate reducers became substrate limited. Thus, the presence of sulfate may be the most important factor controlling the formation of methane in this sediment. However, it is still unclear why such high sulfate concentrations and profiles are observed in sediment from Zegvelderbroek. The influx and oxidation of S-rich organic matter and iron sulfide might explain these relatively high concentrations [40] and its seasonal dynamics might also explain the parallel displacement of the observed concentration profiles. The observed potential consumption rates for 13C-acetate, sulfate and nitrate were comparable to rates determined in summer 1994 and spring 1995. The potential consumption rates for 13C-acetate, sulfate and nitrate determined in autumn 1995 and winter 1995/1996 were 4–6 times lower, suggesting a seasonal pattern (Scholten, unpublished).

Enumerations showed that the dominant acetate-utilising microorganisms were methanogens (2×108 cells cm−3 sediment) and sulfate reducers (2×108 cells cm−3 sediment). The most abundant acetate-utilising methanogen and sulfate reducer were isolated from the highest dilution step of the MPN series and were characterised physiologically and phylogenetically [28]. On the basis of their partial 16S rRNA sequences it became clear that the methanogen and sulfate reducer were closely related to Methanosaeta concilii and Desulfotomaculum acetoxidans, respectively. Both the methanogen (strain AMPB-Zg) and sulfate reducer (strain ASRB-Zg) displayed Vmax, Km and maximum growth rate (μm) values [28] similar to and even slightly lower than described for other acetate-utilising methanogens and sulfate reducers [11,12,33]. From the Vmax and Km values for acetate, determined in resting cell suspensions, it became clear that the most abundant acetate-degrading sulfate reducer strain ASRB-Zg had slightly better kinetic properties than the most abundant acetate-degrading methanogen strain-AMPB. This, in combination with the low acetate concentrations compared to the Km values, explains why acetate-utilising sulfate reducers (in the presence of sufficient sulfate) were able to outcompete acetate-utilising methanogens for the available acetate in the sediment incubations. Besides, strain ASRB-Zg is a generalist and it is possible that acetate degradation is not the only activity of the strain in the sediment. The ability to use other substrates besides acetate can give strain ASRB-Zg a competitive advantage over strain AMPB-Zg (a specialist) when sufficient sulfate is present [28]. At low sulfate concentrations versatile acetate-degrading sulfate reducers may prefer other substrates than acetate [12]. Unfortunately, no information is available on how mixed substrate utilisation may affect the competition between strain AMPB-Zg and strain ASRB-Zg.

The acetate-utilising microorganisms in the sediment were quantified by the MPN method in liquid media. Our MPN counts were similar or 1–2 orders of magnitude higher than the numbers obtained by other investigators in rice field soil, lake and marine sediments for acetate-utilising methanogens and sulfate reducers [41–44]. In these studies the longest incubation period for the MPN counts was 3 months and relatively high temperatures (28–35°C). To obtain true numbers, our incubations needed up to 9 months of incubation after which the results were unaffected by another 3 months of incubation. For example, after 4–5 months of incubation our MPN counts were 2–3 orders of magnitude lower than the final numbers. Bak and Pfennig [45] already mentioned that prolonged incubation times might positively influence the counting efficiency. A few parameters that can be identified as being responsible for prolonged incubation times are growth rate and incubation temperature. Normally the type of MPN counts applied here results in an underestimation of the number of cells [46]. However, the estimation of the number of acetate-utilising methanogens and sulfate reducers based on consumption rates of acetate coincided with the numbers obtained with the MPN method. The results obtained with the independent approach of the consumption rate method, despite its errors, add credence to the estimates of active in situ populations of acetate-utilising microorganisms obtained by the MPN method.

Nitrate-reducing bacteria hardly competed with methanogens and sulfate reducers for the available acetate in the presence of nitrate (Table 2). Nitrate reducers (5×105 cells cm−3 sediment) were clearly outnumbered by the methanogens and sulfate reducers. Furthermore, the pore water profiles showed that nitrate concentrations were below 1 μM. These factors may explain why acetate-utilising nitrate reducers played a minor role in the degradation of acetate in the sediment. Although acetate was consumed, our model was not able to describe this consumption on the basis of competition between acetate-utilising nitrate-reducing bacteria and other microorganisms (Fig. 3C). The low number of denitrifiers in our MPN counts may be a result of the sediment mixing of the upper 10 cm prior to the enumeration and sediment incubation studies. They may be found in large number in the upper part of the sediment where oxygen and nitrate are present at higher concentrations. In this case the competition with the other functional groups may be on kinetic grounds. However, it seemed that in the presence of nitrate other interactions besides competition for acetate played a role in the sediment.

The calculation of electron recovery indicated that, in addition to acetate, nitrate reducers (Table 4) used other electron donors. We observed that part of the nitrate reduction was coupled to the oxidation of reduced sulfur compounds (formation of sulfate) rather than to the oxidation of acetate (Figs. 2C and 3C). Nitrate reduction coupled to the oxidation of reduced sulfur compounds has been reported in studies of natural environments [47–49]. We isolated an acetate-utilising nitrate reducer (strain ANRB-Zg) that was capable of oxidising thiosulfate to sulfate in the presence of acetate [28]. Furthermore, we observed the oxidation of sulfide to sulfate in MPN incubations (106 cells cm−3 sediment) with medium containing sulfide and NO3 (results not shown). This strongly suggests that sulfur-oxidising nitrate reducers were present in the sediment and are most likely responsible for the sulfate accumulation in the presence of nitrate (Figs. 2C and 3C). Chemolithoautotrophic denitrifiers may compete with heterotrophic (acetate-utilising) denitrifiers in the sediment for the available nitrate and this might have led, in combination with possible mixed substrate utilisation, to the observed incomplete acetate and sulfate balances. The model, although it included chemolithoautotrophic denitrification, was also unable to describe acetate and sulfate dynamics satisfactorily at high nitrate concentrations (Figs. 2C and 3C). Only at those conditions, model simulations were significantly different (i.e. differing more than 2 times the standard error) from experimental results. Our experimental and modelling results suggest that the role of chemolithoautotrophic denitrification and mixed substrate utilisation needs more attention to describe substrate dynamics in the presence of available nitrate.

Table 4.  Consumed 13C-acetate, sulfate and nitrate and produced 13C-methane and sulfate (μmol l−1 h−1) obtained from incubations of freshwater sediment samples for 6 h (September 1995) or 7 h (April 1996) at 17°C
  1. Electron recovery calculated from the amounts of 13C-acetate, sulfate, nitrate and 13C-CH4 consumed or formed at the end of the incubations (see Figs. 2 and 3 for values). The standard errors (1 S.E.M.) of the consumed/produced compounds are shown in parentheses.

  2. aMethanogenesis.

  3. bSulfate reduction.

  4. cNitrate reduction.

IncubationConsumedProduced 13C-CH4Electron recovery (%)
 13C-acetateSO42−NO3 masrbnrctotal
September sample
Acetate131 (18)20 (17)0 (0)82 (12)725077
Unsupplemented control0 (0)14 (4)0 (0)0 (0)    
Acetate+SO42−124 (23)83 (18)0 (0)43 (8)4023062
Acetate omitted control0 (0)55 (13)0 (0)0 (0)    
Acetate+NO3100 (27)0 (0)180 (12)1 (5)111213
Acetate omitted control0 (0)0 (0)163 (34)0 (0)    
April sample
Acetate+SO42−126 (12)144 (48)0 (0)34 (1)30940124
Acetate omitted control0 (0)26 (28)0 (0)0 (0)    
Acetate+NO365 (6)99 (78)215 (25)0 (0)060565
Acetate omitted control0 (0)60 (47)210 (27)0 (0)    

In addition to substrate competition, some processes may be inhibited by the accumulation of products. Possible inhibiting products are sulfide, nitrite, NO and N2O [13,15,16]. However, within the sediments sulfide concentrations (0.2–0.5 mM) were always below the concentrations inhibitory of sulfate reduction and methanogenesis (>1.5 mM) [50,51]. Thus this inhibition will be relatively unimportant in the freshwater sediments investigated. However, this might not have been the case for products of nitrate reduction. Brunet and Garcia-Gil [52] mentioned that the initial concentration of free sulfide determines the type of nitrate reduction. At very low concentrations of free sulfide (<50 μM) nitrate was reduced to N2 whereas at high sulfide (>0.3 mM) incomplete reduction to nitrite, NO and N2O took place [52,53]. We were not able to detect the formation of nitrite, NO and N2O in our experiments (detection limits: 1 μM for nitrite and 0.5 mM for NO and N2O, respectively) but sulfide concentrations in the incubations were high enough to affect the type of nitrate reduction. Furthermore, methane production was inhibited in our incubations with nitrate. This inhibition could not be explained by competition for acetate (as acetate concentrations were not limiting (Fig. 3C). Earlier studies have shown that nitrite, NO and N2O inhibited methanogenesis [13,15,16]. Klüber and Conrad [18] showed that NO at concentrations as low as 1–2 μM inhibited acetate-dependent methanogenesis completely. If we take into account that the sulfide concentrations were between 0.1 and 0.5 mM in the incubations, inhibition of methanogenesis in the incubations with nitrate might be explained by the accumulation of low concentrations of NO (<0.2 mM) that were below the detection limit. Therefore, we suggest that inhibition of methanogenesis by NO has to be incorporated in the model in order to describe the influence of nitrate on methanogenesis correctly.

In conclusion, the combination of experiments and modelling proved to be very helpful to understand the fate of acetate under different redox conditions in freshwater sediment. This combination revealed that the competition for acetate between methanogens and sulfate reducers is the main cause of inhibition of methanogenesis in the sediment studied. Nitrate-reducing bacteria hardly competed with methanogens and sulfate reducers for the available acetate. The low number of nitrate reducers may explain why acetate-utilising nitrate reducers played a minor role in the degradation of acetate in the sediment. Furthermore, a significant part of the nitrate consumption was coupled to electron donors other than acetate, most likely reduced sulfur compounds. To fully understand these processes more work has to be done on the organisms involved in the conversion of alternative electron donors in the presence of nitrate.

Acknowledgements

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Methods
  5. 3Results
  6. 4Discussion
  7. Acknowledgements
  8. References

This study was subsidised by grants from the Netherlands Organisation for Scientific Research (NWO) and the Dutch National Research Program on Global Air Pollution and Climate Change.

References

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Methods
  5. 3Results
  6. 4Discussion
  7. Acknowledgements
  8. References
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