HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS		QUICK SEARCH:   [advanced] Author: Keyword(s): Year:  Vol:  Page:

This Article Abstract Figures Only Full Text (PDF) Free Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in ISI Web of Science Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via ISI Web of Science (9) Citing Articles via Google Scholar Google Scholar Articles by Bergman, I. Articles by Nilsson, M. Search for Related Content PubMed Articles by Bergman, I. Articles by Nilsson, M. GeoRef GeoRef Citation Agricola Articles by Bergman, I. Articles by Nilsson, M.

DIVISION S-3-SOIL BIOLOGY & BIOCHEMISTRY

Degradation of ¹³C–U–Glucose in Sphagnum majus Litter

Responses to Redox, pH, and Temperature

^a Dep. of Forest Ecology, Swedish Univ. of Agric. Sci., S-901 83 Ume, Sweden
^b Dep. of Physical Chemistry, Univ. of Ume, S-901 87 Ume, Sweden
^c Pacific Forestry Centre, Natural Resources Canada, 506 West Burnside Rd., Victoria, British Columbia, Canada V8Z 1M5

inger.bergman{at}sek.slu.se

	ABSTRACT

TOP ABSTRACT INTRODUCTION Materials and methods Results Discussion Conclusions REFERENCES

 
We studied the utilization of ¹³C–U–glucose by the microbial community in shallow Sphagnum majus (Russ.) C. Jens. ssp. norvegicum Flatb. litter and its regulation by pH, temperature, and redox conditions. The transformation of ¹³C–glucose was monitored by solution- and solid-state ¹³C–nuclear magnetic resonance (NMR) spectroscopy. The aerobic microbial community used the glucose C for respiration and, to a lesser degree, for storage as mannitol, triglycerides, and polysaccharides. Under both aerobic and anaerobic conditions, the allocation of glucose C for storage was greater at pH 6.8 than at 4.3; however, the amount of C used for building new biomass was the same at both pH settings. At 15°C, 15 to 18% of the utilized C under aerobic conditions was found in new microbial biomass: less than the previously reported values of 40 to 72%. This indicates that peat soils may promote significantly different microbial growth patterns from other minerogenic and moor humus soils. The production of mannitol and triglycerides suggests that fungi dominated the microbial community and utilized the glucose under aerobic conditions. Using a combination of solid and liquid NMR techniques we were able, for the first time, to follow the anaerobic pathways of glucose degradation in a natural soil sample. The anaerobic microbial community produced mainly volatile fatty acids (VFA), ethanol, and CO₂ from the added glucose, and only minor amounts were converted to methane, storage C, and new microbial biomass. Nuclear magnetic resonance spectroscopy allows nondestructive assays of metabolic events and, therefore, was shown to be an excellent tool for studying the microbial utilization of ¹³C–glucose in peat.

Abbreviations: ¹³C–NMR, carbon-13 nuclear magnetic resonance • CP, cross polarization • CPMAS, cross-polarization magic-angle spinning • FAC, fatty acyl carbon • Glc, glucose • ni, not identified • NMR, nuclear magnetic resonance • PBHB, poly-ßß-OH-butyrate • SSB, spinning sidebands • VFA, volatile fatty acid

	INTRODUCTION

TOP ABSTRACT INTRODUCTION Materials and methods Results Discussion Conclusions REFERENCES

 
THE PEATLANDS OF THE WORLD are highly concentrated C sinks, so the global C dynamics of these peatlands are of major importance for global C circulation. In peatlands, C accumulates because of the high water content and because the annual input of dead plant material exceeds annual breakdown. Generally, the organic matter in soils consists of fresh plant litter that has not yet been altered by microorganisms, old plant C resistant to microbial attack, and microbially transformed C compounds. Transformed C, such as microbial polysaccharides, may be further decomposed, whereas lipids are more recalcitrant and accumulate during microbial decomposition of the organic matter. Selective loss of carbohydrate C (O-alkyl) and net accumulation of lipid C (CH₂)_n has been reported to occur during decomposition of both forest litter and peat (Baldock and Preston 1995). Studies using NMR techniques have also revealed a lower aromatic content than previously believed, while lipids seem to be the major fraction accumulated (Baldock et al., 1992).

Glucose is the most commonly used model substrate to study both aerobic and anaerobic decomposition processes, as it is a major constituent of organic matter. Glucose made available to the microorganisms can be utilized for energy production, storage, or for new microbial biomass. Energy production under aerobic conditions results in the release of CO₂, whereas both methane and CO₂ are produced under anaerobic conditions. When microbial activity is not restricted by energy or C limitations, the cells are able to produce storage compounds within the cells. Storage compounds accumulated in different microorganisms include mannitol, poly-ßß-hydroxy butyrate (PBHB), glycogen, and various lipids (Martin et al., 1985; Zelles and Bai, 1993; Zelles et al., 1995; Cavigelli et al., 1995). Mannitol and unsaturated lipids are typical storage compounds for fungi (Harwood and Russell, 1984; Richards, 1987; Martin et al., 1988), whereas glycogen and PBHB are typical storage compounds for bacteria, although glycogen is also produced by many eucaryotic organisms (Stanier et al., 1986; Vanderhart et al., 1995). The reported efficiencies, i.e., the proportion between assimilated and respired C, for microbial growth after glucose additions to soils lie between 40 and 72% (Killham et al., 1993; Bremer and Van Kessel, 1990; Bremer and Kuikman, 1994; Tsai et al., 1997).

The aerobic microbial decomposition of ¹³C–glucose in soils has also been studied using the technique of ¹³C–CPMAS (cross polarization magic angle spinning) NMR spectroscopy for solid-state samples (Baldock et al., 1989, 1990a; Golchin et al., 1996; Webster et al., 1997). Solid-state CPMAS analysis of the microbial utilization of C from ¹³C glucose showed that ¹³C was mainly allocated to the O-alkyl, alkyl, acetyl, and carbonyl fractions of soils (Baldock et al., 1989; Golchin et al., 1996; Webster et al., 1997). Baldock et al. (1990a,b) also showed there were differences in the chemical structure of the C synthesized from fungal and bacterial cultures, finding a larger proportion of O-alkyl C and a smaller proportion of alkyl C associated with the fungi. Solution ¹³C–NMR spectroscopy has also been used to study anaerobic metabolic pathways in pure cultures of bacteria (Roberts et al., 1987; Houwen et al., 1991) and ectomycorrhizal fungi (Martin, 1991).

In solution ¹³C–NMR, mobile C atoms are determined in a quantitative manner because the peaks are well resolved. Also, these high resolution spectra make it possible to identify the carbons and assign them to certain compounds. The spectra from solid-state ¹³C–NMR have lower resolution than solution NMR spectra, along with associated problems for quantification (Preston, 1996). On the other hand, solid state NMR spectra give information about immobile carbons in the soil that cannot be derived from solution ¹³C–NMR spectra. In this study we have used peatland soil as a model system for studying microbially mediated decomposition processes. Our objectives were to study how the utilization of ¹³C–U–glucose by the microbial community in shallow S. majus litter is regulated by pH, temperature, and redox conditions.

Studies on the regulation of terminal C mineralization (i.e., into methane and CO₂) are presented elsewhere (Bergman et al., 1999). In this paper we focus on the transformation of glucose into storage C, VFAs, and new cell biomass. The transformation of ¹³C glucose added to S. majus litter in an aqueous slurry was monitored by solution- and solid-state ¹³C–NMR spectroscopy, the production of methane and CO₂ by gas chromatography, and the fractions of ¹³C-labeled CO₂ and CH₄ were obtained by mass spectrometry.

	Materials and methods

TOP ABSTRACT INTRODUCTION Materials and methods Results Discussion Conclusions REFERENCES

 
Study Site and Sampling
Samples of litter from Sphagnum majus plant material were collected in September 1996 from approximately 10 to 15 cm below the moss surface, just below the green parts of the mosses. The site was at Stormyran, a flark level bog (Eurola et al., 1984) in an acid–mixed mire, pH 3.5 to 4.3, 10 km from the coast, situated 15 km south of Ume, Sweden (63° 44' N, 20° 06' E, 35 m altitude), with a mean air temperature of approximately +15°C in the growing season (for site description see Mikkelä et al., 1995). The sampling site comprises a wet carpet dominated by S. majus and is largely covered by the vascular plant Rhynchospora alba L., with the litter almost permanently waterlogged throughout the growing season.

Litter was collected from an area of approximately 0.5 m², put in a plastic bag, filled with pore water from the site, sealed, and immediately brought to the laboratory for preparation. The incubations were started within 10 h of sample collection.

Experimental Conditions
Samples were mixed in a blender under a continuous flow of N₂, and subsamples (4–5 g wet weight) were transferred into sterile incubation flasks (58 mL) that were previously filled with sterile distilled water or phosphate buffer (10 mM, pH 6.8). A sterile solution of [U–¹³C]–glucose (99.9%; 10 mM; supplied by Larodan Fine Chemicals AB, Malmö, Sweden) was added to all flasks except those which were incubated without substrate addition. The final water volume in each flask was 10 mL. The samples incubated under aerobic conditions were closed with a cotton plug at atmospheric pressure and the anaerobic samples were sealed with a butyl rubber stopper. Aerobic CO₂ production rates were measured by replacing the cotton plug with a butyl rubber stopper and thereafter sampling the accumulated gas after 3, 6, and 9 h. Finally, the rubber stopper was replaced by the cotton plug. For the aerobic samples this procedure was repeated for eight occasions during the incubation period. The anaerobic set of samples was evacuated and refilled with N₂ to 0.2 MPa in three consecutive cycles. All samples were incubated at 15 or 25°C in darkness, with slow shaking. Prior to the first gas analysis, the samples were incubated for 48 h at the predetermined conditions. The amount of accumulated CH₄ or CO₂ was measured by collecting gas samples from the headspace of the flasks during the experimental period using an airtight syringe.

All samples without added glucose (Number 1–8, Table 1) were run in duplicate. Each experimental combination with labeled glucose supplement (Number 9–16, Table 1) was carried out with six replicates. The biological activity was terminated in one of each of the three replicates of the glucose-amended treatments after two, three, and four weeks, by adding 0.1% NaN₃ (v/v). The three terminated samples from each treatment provided a time series of glucose degradation for that treatment (a total of 24 samples) and were analyzed by solution-state ¹³C–NMR. The remaining three samples for each treatment and the duplicate samples that were not amended with glucose were terminated after four weeks of incubation.

Gas Analyses
Methane and CO₂ were determined using a Perkin Elmer (Norwalk, CT) Sigma II gas chromatograph equipped with a flame ionization detector and a column packed with Porapak Q 80/100 mesh (J & W Scientific, Folsom, CA). For the analysis of CO₂, a methanizer for reducing CO₂ to methane was connected prior to the flame ionization detector. Nitrogen was used as carrier gas at a flow rate of 30 mL min^-1. Injector, oven, and detector temperatures were set at 35, 40, and 350°C, respectively. The detector response was linear in the range 1.7 to >3000 ppm. Repeated injections of methane standards (100–3000 ppm) using a 0.5-mL gas-tight glass syringe resulted in a standard deviation of <3% of the injected concentration.

Methane labeled with ¹³C was measured by mass spectrometry (Europa Scientific 20-20 stable isotope analyzer with an ANCA-NT system gas purification module, Crewe, UK). Carbon dioxide labeled with ¹³C was not measured because of instrument problems. Instead, the ¹³C–CO₂ was calculated as the difference between the amount of CO₂ produced after addition of ¹³C–glucose and the amount of CO₂ produced without glucose addition.

The amount of CH₄ dissolved in water decreases with increasing temperature, but lies between 3 to 5% of the total CH₄ content within the temperature range used (Merck Index, 1989) and was, therefore, ignored in the calculations. The variation of solubility of CO₂ in water with pH and temperature was taken into account in the calculations for the rate of CO₂ production using Henry's Law constants (Stumm and Morgan, 1996).

In this study we report the amount of methane and CO₂ accumulated in the samples used for ¹³C–NMR spectroscopy, after 2, 3, and 4 wk of incubation. The amount of accumulated CO₂ produced aerobically was extrapolated linearly across time intervals from the measured rates of CO₂ production.

Solution-State ¹³C–NMR
Carbon-13 solution spectra of the peat slurries were obtained at 125.77 MHz on a Bruker AMX/2-500 MHz NMR spectrometer using 10-mm-o.d. tubes at an operating temperature of 25.0°C in a Bruker broadband probe. The acquisition conditions were: acquisition time, 0.293 s; relaxation delay, 1.2 s; 16K datapoints; 90° pulse; 23.8 µs spectral width; 27933 Hz; and 600, 1200, or 2400 accumulated transients. To eliminate sample heating and optimize nuclear Overhauser effect (nOe) signal enhancement, WALTZ-16 ¹H decoupling was used in a bilevel decoupling scheme.

Two individual samples were used for the saturation factor determinations, both correcting for the nuclear Overhauser effect and T₁ effects: one pretreated anaerobically for two weeks at pH 6.8 and 25°C, and the other for two weeks at pH 4.3. The chemical shifts were referenced to DSS (2,2-dimethyl-2-sila-pentane-5-sulfonic acid) using [ßß-C1–¹³C–]–glucose as a secondary reference assigned to 97.0 ppm.

Peak Assignments
Some resonances were assigned according to published data (Rutar et al., 1977; Bengsch et al., 1986; Fan, 1996). The resonances for several metabolites were assigned by analyzing samples of pure compounds in an aqueous solution. For unsaturated lipids, the oil seeds of flax, Linum usitatissimum L., were used. The chemical shift of mannitol from our spectra deviated from those found experimentally in pure solution. We found that the deviation of the chemical shifts of pure mannitol in an aqueous solution (100 mM) from the resonances in the samples was due to complex binding of the mannitol with Fe²⁺ (Lundberg et al., personal communication, 1998). The chemical shifts for all compounds identified by solution ¹³C–NMR are presented in Table 2 . There were no signals from the native ¹³C in the solution ¹³C–NMR spectra (data not shown) of control samples; therefore, all resonances obtained were assigned as C from the added ¹³C–glucose. The spectrum for 10 mM ¹³C–glucose (Fig. 1a) shows the intensities of the resonances assigned to [U–¹³C]–glucose prior to degradation.

View larger version (18K): [in this window] [in a new window]   Fig. 1 Solution NMR spectra of aerobically degraded 13C glucose in samples incubated for four weeks. Assignments are presented in Table 2 except for PS (Fig. 1D) which refers to resonance from polysaccharide C. A, unsupplemented sample with addition of 10 mM 13C glucose just before the NMR analysis. B, 15°C and pH 4.3; C, 25°C and pH 4.3; D, 15°C and pH 6.8 E, 25°C and pH 6.8. (=) and (= =) refers to monounsaturated C and polyunsaturated carbons, respectively. Note that the scale of relative intensity is magnified in fig. 1E and 1D

Because of limitations in the amount of instrument time available, it was not possible to analyze all the samples by this method. We analyzed all of the 4-wk samples with glucose supplements, and four of the 4-wk samples without glucose supplements (only those incubated at 25°C). Spectra were also obtained for the ¹³C–glucose, as well as for one time-series from the samples incubated anaerobically at 15°C and pH 6.8. The samples used for solid-state NMR analysis were composites of the two replicates.

Unlike the solution NMR, the solid-state spectra included the background of the natural abundance ¹³C in the peat. We consequently used the approach of generating difference spectra with respect to the unsupplemented samples to examine changes in the added ¹³C–glucose. Spectra of the control samples incubated without glucose were all very similar. Three of these samples were used to prepare a composite sample, whose spectrum was subtracted from the spectrum of each sample supplemented with ¹³C–glucose with the following procedure. Using the dual-display mode in the Bruker software, first the baselines were aligned, and then the height of the control spectrum was decreased to match the peak heights of the sharp maximum at 105 ppm. This was chosen after careful examination of both the solution and solid-state spectra did not indicate formation of ¹³C products with a peak at 105 ppm. Anomeric signals for residual ¹³C–glucose occur at 92 and 96 ppm, and for the new microbial ¹³C polysaccharide at 90 to 103 ppm. The general validity of this procedure was supported by the absence of any negative peaks after subtraction, and by a general consistency with the metabolites observed by solution ¹³C–NMR. Also, the S/N of the difference spectra was obviously lower for the two incubation conditions with the greatest loss of ¹³C as CO₂ (aerobic, pH 6.8, 15, and 25° C, Fig. 5c, 5d) . Area integrals of the difference spectra were analyzed for the following regions: alkyl C (0–50 ppm), O-alkyl C (50–90 ppm), di-O-alkyl C (90–110 ppm), aromatic (110–165 ppm), and carbonyl carbons (165–200 ppm), and expressed as percentage of total intensity.

View larger version (14K): [in this window] [in a new window]   Fig. 5 Solid state NMR difference spectra of 13C–glucose supplemented samples incubated aerobically for four weeks at A, 15°C pH 4.3; B, 25°C and pH 4.3; C, 15°C and pH 6.8; D, 25°C and pH 6.8

View larger version (14K): [in this window] [in a new window]   Fig. 4 Solid state NMR spectra of A, composite sample without addition of glucose; B, pure 13C–glucose; C, normal spectrum, and D, difference spectrum (i.e. A subtracted from spectra C) of samples supplemented with glucose and incubated aerobically for four weeks at 25°C and pH 6.8. Spinning side bands of carboxyl C and 13C–glucose C are denoted as xx and x, respectively

View larger version (21K): [in this window] [in a new window]   Fig. 6 Carbon types identified by solid state NMR produced from the degradation of 13C–glucose in samples incubated under aerobic conditions at different pH and temperature conditions

The assignments of groups of C from the solid-state spectra were alkyl carbons (0–50 ppm), O-alkyl C (50–90 ppm), di-O-alkyl C (90–110 ppm), and carbonyl carbons (170–182 ppm).

	Results

TOP ABSTRACT INTRODUCTION Materials and methods Results Discussion Conclusions REFERENCES

 
Aerobic Conditions, Solution State ¹³C–NMR
For both incubation temperatures at the in situ pH of 4.3, (Fig. 1b and 1c), most of the soluble ¹³C remaining was still found as glucose (61–97 ppm) after four weeks of incubation, with a small amount of fatty acyl chain carbons (FAC; 14–34 ppm). In contrast, the spectrum at pH 6.8 and 15°C (Fig. 1d) revealed the presence of several new carbons in addition to those attributable to glucose after four weeks of incubation. The anomeric carbons of glucose were still present at 93 and 97 ppm, and the three large peaks at 66.1, 71.6, and 74.1 ppm were identified as mannitol constituents. Resonances for FAC were found between 14 to 34 ppm, with the largest peak at 30 ppm and small resonances at 129 ppm for C in double bonds and at 175 ppm for the carbonyl carbons (Fig. 1d). Broad, weaker resonances were also found underlying the O-alkyl and di-O-alkyl regions. At 25°C and pH 6.8, no glucose remained and broad peaks were found in the region between 60 to 84 ppm and around 100 ppm after four weeks incubation. These broad peaks were assigned as unidentified O-alkyl (60–80 ppm) and di-O-alkyl carbons (100 ppm) and were most likely due to microbial polysaccharides. Figure 2 shows solution ¹³C–NMR spectra of the time course for degradation of ¹³C–glucose at 15°C and pH 6.8. The resonances from glucose carbons decreased, and those for mannitol (66.1, 71.6, and 74.1 ppm), FAC (14–34 ppm), and carbonyl C (175 ppm) increased between two and four weeks of incubation (Fig. 2a–c).

View larger version (16K): [in this window] [in a new window]   Fig. 2 Solution NMR spectra of aerobically degraded 13C–glucose in samples incubated at 15°C and pH 6.8 after A, two weeks; B, three weeks; and C, four weeks. Assignments are presented in Table 2. (=) refers to monounsaturated carbons. Peak assignments 1 through 6 in the alkyl region between 14.3 ppm and 33.5 ppm refer to FA1 through FA6 in Table 2. Note that the scale of relative intensity is magnified in Fig. 2C

View larger version (24K): [in this window] [in a new window]   Fig. 3 Carbon types identified by solution NMR and CO2 produced from the degradation of 13C–glucose in the samples incubated aerobically for four weeks at A, 15°C and pH = 4.3; B, 25°C and pH = 4.3; C, 15°C and pH = 6.8; D, 25°C and pH = 6.8

Aerobic Conditions, Solid-State NMR
Solid-state ¹³C–NMR spectra of the samples incubated without ¹³C–glucose were very similar (data not shown). Three of these samples were used to prepare a composite sample (Fig. 4a) that was used to generate the difference spectra. The spectrum is typical for poorly humified sphagnum peat with the main resonances for O-alkyl (50–90 ppm) and di-O-alkyl (90–110 ppm) carbons, and smaller features for alkyl (0–50 ppm) and carbonyl C (170–182 ppm) (Holmgren et al., 1990, Nordén et al., 1992). There was little difference from a spectrum obtained from fresh sphagnum plant material (Dudley et al., 1990). The spectrum of pure ¹³C–glucose has peaks for anomeric C1 at 92 and 96 ppm, and from the other carbons at 61, 63, and 71 ppm (Fig. 4b). The peaks are broadened due to ¹³C–¹³C coupling (Baldock et al., 1989). Figure 4d shows an example of a difference spectrum generated by subtraction of the composite control sample from the S. majus litter incubated with ¹³C glucose at 25°C and pH 6.8 (Fig. 4c).

The difference spectra of the 4-wk samples incubated at pH 4.3 (Fig. 5a,b) show that the main peaks are from unaltered ¹³C–glucose, in agreement with the results from the solution NMR spectra (Fig. 1b,c). For samples incubated at pH 4.3 the intensities from alkyl carbons and carbonyl carbons were higher at 25°C than at 15°C (Fig. 5a,b). The difference spectra from the samples incubated under aerobic conditions at pH 6.8 (Fig. 5c,d) show larger peaks from both alkyl carbons (7–41 ppm) and carboxyl carbons (170–183 ppm) compared with the samples incubated at pH 4.3. In addition, no remaining ¹³C–glucose was detected in the samples incubated at pH 6.8, based on the absence of signal from the anomeric carbons at 92 and 96 ppm. Transformation of the ¹³C–glucose into other carbohydrate structures was indicated by the peaks for O-alkyl and di-O-alkyl C at 72 ppm and 102 ppm (Fig. 5c,d). At pH 6.8 under aerobic conditions, the resonances from alkyl C and carboxylic carbons were somewhat lower at 25°C than at 15°C for the incubated samples (Fig. 5c,d). For all the samples incubated aerobically, the major resonances [underscored] in the alkyl region were consistent with production of acetyl C (R-CH₂-COO^-) or methyl C (CH₃-R), methylene C at the C2 position (CH₃-CH₂) at 21 ppm and polymethylene carbons (-CH₂-CH₂-CH₂-) of long aliphatic chains at 30 ppm (Fig. 5).

The quantitative results from the solid-state NMR spectra after four weeks of incubation (Fig. 6 and Table 3) represent ¹³C from the glucose addition found in a less mobile or insoluble form; e.g., as in cell wall constituents, or in organic material adsorbed to peat particles that were not detected by solution NMR. For the aerobic samples, the ¹³C compounds other than glucose detected in the CPMAS spectra are thought to represent the fraction of C mainly used for new cell biomass. For the samples incubated under aerobic conditions <10% of the total ¹³C added was allocated to new cell biomass, except for the sample incubated at 15°C and pH 6.8, in which 16% was found as cell biomass (Table 3). At pH 4.3 only FACs were identified in the solid fraction. At pH 6.8 both new carbohydrate C and FACs were allocated to the cell fraction (Table 3). However, the amounts of FACs in the cell fraction were larger at pH 4.3 than in the samples incubated at pH 6.8 (Fig. 6). The abundance of FACs produced was greater at 25°C than at 15°C, and the proportional difference was similar at both pH settings (Fig. 6). In contrast, at pH 6.8, there was approximately 10% less new carbohydrate C produced at 25°C than at 15°C (Table 3).

Aerobic Conditions, Carbon Dioxide Production
For the aerobic incubations, the production of CO₂ was highest from the samples incubated at pH 6.8 (P < 0.01; Bergman et al., 1999). At 15°C the amount of CO₂ produced increased until termination of the experiment after four weeks (Fig. 3c), whereas at 25°C the amount decreased between the third and fourth week of incubation (Fig. 3d). From the samples incubated for four weeks at 15 and 25°C, 50 and 77% of the added ¹³C was recovered as CO₂, respectively (Table 3). At pH 4.3, the production of CO₂ increased with both temperature (P < 0.05; Bergman et al., 1999) and time (Fig. 5a,b). After four weeks of incubation at pH 4.3, 21 and 27% of the added glucose was transformed to CO₂ at 15 and 25°C, respectively (Table 3).

Anaerobic Conditions, Solution-State ¹³C–NMR
Addition of ¹³C–glucose to unamended samples incubated anaerobically for four weeks just prior to performing the solution NMR analysis resulted in acquisition of a pure glucose spectrum (Fig. 7a) . After four weeks of incubation at pH 4.3 and 15°C, resonances observed in the glucose-amended peat were dominated by glucose (Fig. 7b), whereas the resonances from the sample incubated at 25°C were attributable solely to alkyl and carboxylic carbons (Fig. 7c). After four weeks of incubation at pH 6.8, the spectra were dominated by resonances from alkyl and carboxylic carbons with some additional broad resonances from unidentified polymeric carbons in the region between 70 and 100 ppm (Fig. 7d,e). Glucose was only found in the two-week sample incubated at 15°C, and not at all after three or four weeks (Fig. 8a) . At both temperatures, the resonances from alkyl (10–64 ppm) and carboxylic carbons (180 and 183 ppm) dominated and increased between two and three weeks of incubation (Fig. 8 and 9) . Resonances from the unidentified polysaccharide carbons (61 and 71 ppm) were equal after two and four weeks of incubation at either temperature, but greater in the spectra from the samples incubated at 15°C (Fig. 8) than in those incubated at 25°C (Fig. 9).

View larger version (14K): [in this window] [in a new window]   Fig. 7 Solution state NMR spectra of anaerobically degraded 13C–glucose in samples after four weeks of incubation. Assignments are presented in Table 2. A, control sample with addition of 10 mM 13C–glucose just before the NMR analysis; B, 15°C and pH 4.3; C, 25°C and pH 4.3; D, 15°C and pH 6.8; E, 25°C and pH 6.8. Note that the scale of relative intensity is magnified in E and D

View larger version (18K): [in this window] [in a new window]   Fig. 8 Solution NMR spectra of anaerobically degraded 13C glucose in samples incubated at 15°C and pH 6.8 after A, two weeks; B, three weeks; and C, four weeks. Assignments are presented in Table 2. Note that the scale of relative intensity is magnified in C

View larger version (25K): [in this window] [in a new window]   Fig. 9 Solution state NMR spectra of anaerobically degraded 13C glucose in samples incubated at 25°C and pH 6.8 for A, two weeks; B, three weeks; and C, four weeks. Assignments are shown in Table 2. Note that the scale of relative intensity is magnified in C

View larger version (27K): [in this window] [in a new window]   Fig. 10 Carbon types identified by solution NMR or gas analysis produced from the degradation of 13C–glucose in samples incubated anaerobically at A, 15°C and pH 4.3; B, 25°C and pH 4.3; C, 15°C and pH 6.8; D, 25°C and pH 6.8. Note that the scale is magnified for some bars. Very low or zero amounts are written in the Figure for the specific compound

Anaerobic Conditions, Solid-State NMR
At the in situ pH of 4.3, the glucose was degraded to a very small extent as shown by the CPMAS difference spectra (Fig. 11a,b) , which were dominated by the peaks of glucose after four weeks of incubation. However, the resonances from alkyl C and carbonyl carbons were considerably higher in the sample incubated at 25°C (Fig. 11b). The added glucose was completely degraded after three weeks of anaerobic incubation at 15°C and pH 6.8 (Fig. 12b) . After two weeks of incubation, the major resonances were still from unaltered glucose (63–97 ppm), but substantial intensity was also seen from alkyl (11–40 ppm) and carbonyl carbons (183 ppm) (Fig. 12a). After three and four weeks at 15°C and pH 6.8 (Fig. 12b,c), the peaks in the O- and di-O-alkyl region (71 and 105 ppm) were from a carbohydrate other than glucose, corresponding to 3.7 and 4.6% of the added ¹³C, respectively. The alkyl and carbonyl carbons produced in samples incubated anaerobically were identified as constituents of VFAs (Fig. 11 and 12).

View larger version (13K): [in this window] [in a new window]   Fig. 11 Solid state NMR difference spectra of 13C–glucose supplemented samples incubated under anaerobic conditions four weeks at A, 15°C and pH 4.3; B, 25°C and pH 4.3; C, 15°C and pH 6.8; D, 25°C and pH 6.8. Spinning side bands of 13C–glucose C are denoted as x

View larger version (14K): [in this window] [in a new window]   Fig. 12 Solid state NMR difference spectra of 13C glucose supplemented samples incubated under anaerobic conditions at 15°C and pH 6.8 for A, two weeks; B, three weeks; and C, four weeks. Spinning side bands of carboxyl C and 13C–glucose C are denoted as xx and x, respectively

Anaerobic Conditions, Methane, and Carbon Dioxide Production
After four weeks of incubation, the amount of CO₂ produced was greatest in the samples incubated at pH 6.8 (P < 0.01; Bergman et al., 1999). At 15°C, 29% of the ¹³C was recovered and at 25°C, 12% was recovered as CO₂ (Table 3). The amount of CO₂ produced at 15°C increased from two to three weeks of incubation but did not change by the fourth week of incubation at 25°C (Fig. 10c,d). At pH 4.3, 1.3 and 7.9% of the added ¹³C was recovered as CO₂ after four weeks of incubation at 15 and 25°C, respectively (Fig. 10b and Table 3). According to mass spectrometric analysis, approximately 60% of the methane produced was enriched with ¹³C. The recovery of ¹³C in methane increased with temperature (P < 0.05, Bergman et al., 1999) but decreased with pH ( ; Bergman et al., 1999). The percentage of ¹³C–CH₄ was three times higher for the samples incubated at pH 4.3 vs. the corresponding sample incubated at pH 6.8 (Table 3).

	Discussion

TOP ABSTRACT INTRODUCTION Materials and methods Results Discussion Conclusions REFERENCES

 
Aerobic Degradation
The active aerobic microbial community used the glucose C for respiration and also, to a lesser degree, for storage and building new microbial biomass (e.g., cell wall components and membranes). This pattern of glucose C utilization by the microorganisms in the S. majus litter varied with pH and temperature. The solution NMR spectra showed ¹³C-compounds in a mobile form, i.e, in solution or oil droplets, whereas ¹³C-compounds allocated to more rigid and insoluble molecular structures, such as cell walls, were revealed from solid state NMR spectra after subtraction of the solid-state NMR difference spectra.

The carbons allocated to the storage fraction of the microbial organisms were found to be constituents of mannitol and FACs. Mannitol, a polyhydroxy alcohol, is the main soluble storage compound of various fungi and actinomycetes (Richards, 1987), including those that are ectomycorrhizal (Martin et al., 1988). The FACs detected probably indicate the occurrence of triglycerides, although the glycerol backbone of the triglycerides was not detected in the solution NMR analysis. Yet, considering the proportion of glycerol carbons that would be expected, in relation to the FACs, the concentration of ¹³C-labeled glycerol would probably be below the detection limit for the solution NMR analysis. Triglycerides are also common storage compounds in fungi, accumulating within the cells as small oil droplets (Harwood and Russell, 1984; Martin et al., 1984). It has also been suggested that previously observed microbial production of triglycerides from degradation of ¹³C–glucose in a forest humus soil occurred through fungal agents (Lundberg et al., personal communication, 1998). In addition to mannitol and FACs, the glucose C also appeared to be allocated into polysaccharide structures, judging by resonances consistent with O-alkyl and di-O-alkyl-carbons in the NMR spectra. This is in accordance with studies of different soils by solid-state NMR analysis, indicating that the carbons from ¹³C–glucose microbially degraded under aerobic conditions are allocated to new carbohydrates in addition to alkyl and carbonyl compounds (Baldock et al., 1989,1990b; Webster et al., 1997).

In the present study, the FAC and carbohydrate C were, at least in part, further allocated by the microorganisms to the solid fraction: i.e., cell biomass. This was concluded from the decrease of these carbons detected by solution NMR between the third and fourth week of incubation, and from the simultaneous increase in their amounts observed by solid state NMR (Fig. 3 and Table 3). In confirmation of the allocation of carbons to the solid fraction, the decrease in soluble storage C was not accompanied by a corresponding increase in CO₂ production. In a study of fine sandy loam, 50% of the alkyl C formed from degradation of ¹³C–glucose was found to be incorporated into mobile polymethylene (CH₂)_n, whereas the other half was incorporated into nonmobile polymethylene chains (Baldock et al., 1989). These mobile and nonmobile forms of polymethylene may have reflected the proportions of lipids allocated in the microbial cells to storage and new microbial biomass, respectively.

The degradation of glucose under aerobic conditions was clearly restricted by acid pH conditions, resulting in a less heterogenous allocation pattern at pH 4.3 than at pH 6.8. Although almost no C was allocated for storage within the cells at pH 4.3, approximately the same amount of C was used for building new biomass at pH 4.3 as at pH 6.8. Although the rate of glucose degradation was lower at pH 4.3, the relative amount of utilized glucose incorporated into new microbial biomass after four weeks was very similar under both pH conditions (15–18%) except at 25°C and pH 6.8. At pH 6.8 and 25°C only 6% of the degraded glucose occurred as new microbial biomass. We believe that the proportion of new cell C at 25°C and pH 6.8 was greater after two weeks of incubation and thereafter decreased due to C limitation. In addition to turnover of the new cell C, the decreased availability of C would be expected to result in a decline of storage C and CO₂ production, as confirmed by the results presented in Fig. 3d. The above conclusions are supported in a study by Tsai et al. (1997). They found that the turnover of biomass C in a sandy loam soil increased during the first three days of incubation after addition of ¹⁴C–glucose, but decreased to the original value after nine days of incubation.

At pH 6.8, the proportion of utilized glucose C allocated for storage was greater than at pH 4.3, owing to the higher turnover rate of glucose at pH 6.8 and to the restricted ability for growth.

Anaerobic Degradation
This is the first study, to our knowledge, that has been able to follow the anaerobic degradation pathways of glucose in a natural peat sample by using a combination of solid and solution NMR techniques. Both solution- and solid-state NMR were used to investigate the effect of temperature and pH on the allocation of glucose C within the anaerobic microbial community in surface S. majus litter. In general, the CPMAS and solution NMR gave consistent results, and the differences were consistent with the analytical limitations with respect to the compounds detectable by the two methods. In addition, ethanol was not observed in the solid-state spectra, as it was lost during freeze-drying of the samples. One exception was observed for the sample incubated at 25°C and pH 4.3 for four weeks, for which there was an obvious discrepancy between solution- and solid-state NMR (Fig. 7c and 11b). The solution NMR spectrum (Fig. 7c) showed no remaining glucose, and all signals were attributable to VFA and ethanol C. The solid-state NMR spectrum (Fig. 11b) showed a large signal from glucose and smaller signals from VFAs. In this study, the solid-state analysis was performed on composite samples of the two replicates, but only one of these samples was analyzed by solution NMR. In this case, there may, therefore, have been a large difference in the degradation of ¹³C–glucose between the two replicates.

Except for the methane production, the overall microbial activity was greater at the higher pH, and the anaerobic microbial community as a whole, including the methanogenic component, was apparently stimulated by the higher temperature. We have previously shown that the temperature response in methane production was highly dependent on the amount of added glucose (Bergman et al., 1998).

Despite the high rate of glucose degradation at pH 6.8, most of the utilized glucose C was accumulated in acetate, propionate, and butyrate (VFA), or respired in CO₂, and only a minor proportion was found in new microbial biomass. The VFAs detected are common degradation products from anaerobic methane-producing microbial communities in these kinds of ecosystems (Krumböck and Conrad, 1991; Westermann, 1994; Schultz and Conrad, 1996; Kotsyurbenko et al., 1996). It is well known that the degradation of VFAs to acetate, H₂, and CO₂ by syntrophic bacteria is endothermic under standard conditions and is only possible within a certain narrow H₂– partial pressure range maintained by exothermic methanogenesis. Above this H₂ partial pressure, fatty acid oxidation is product-inhibited (Dolfing and Tiedje, 1988; Westermann, 1996). In the present study we consequently believe that further oxidation of the accumulated VFAs to substrates for methanogenesis was inhibited by high partial pressures of H₂.

The accumulated VFAs can be considered as temporarily conserved energy that is not available for the microorganisms for use in growth or storage C compounds. In comparison with aerobic microorganisms, the energy available per mole of glucose to the organisms of the anaerobic microbial community is much lower. The reason for this is that the energy released aerobically per mole of glucose is available to just the single organism consuming the glucose, whereas the energy released anaerobically is shared among the members of the microbial community (Schink, 1988; Schink and Friedrich, 1994). However, even though all the conserved energy in the VFAs is available for the methanogens, 96% of the energy released from the VFAs is lost as methane and only 4% is available for the methanogens (Westermann, 1996).

Considering these features, in addition to the nutrient-limited conditions existing in the mire, the utilization of glucose C for storage or new cell C would be expected to be a very slow process within the anaerobic microbial community producing methane. In a study by Sawyer and King (1993), only 20% of added ¹⁴C–glucose was found in the particulate fraction of a marine sediment after one week of anaerobic incubation. Notwithstanding, the authors believed that the relative incorporation of glucose into biomass was probably minimal because of labeled carbons being distributed among the intermediates involved in the various biochemical pathways of the microbial cells.

The total recovery of ¹³C increased during the course of incubation for both the aerobic and anaerobic incubations mainly in the samples where the utilization of glucose was rapid (Fig. 3 and 10). This suggests that some of the ¹³C label may have been found in various metabolites within the cells at concentrations too low for detection by solution NMR spectroscopy. Yet, this further implies that the ¹³C signals of these metabolites occur at separate chemical shifts with no overlap in the solution NMR.

At pH 6.8, large amounts of ethanol were produced, indicating the presence of yeasts. Although some anaerobic bacterial species can carry out alcoholic fermentation, most of the ethanol formed in nature comes from the anaerobic breakdown of glucose and other hexoses by yeasts (Gottschalk, 1979). From the 10- to 20-cm level of a wet ombrotrophic S. majus site from the same mire, four yeast types were isolated according to colony morphology, as were several other fungal species (Nilsson et al., 1992). These four yeast types were very frequent among the fungal isolates. At 25°C, the amounts of ethanol decreased at the end of the incubation period, indicating that the ethanol produced was consumed (Fig. 10d).

Aerobic vs. Anaerobic Degradation
The microbial utilization of glucose C for the production of storage compounds and the formation of new microbial cell biomass was more effective under aerobic conditions than under anaerobic conditions. This was most obvious at pH 6.8, where the amounts of storage compounds decreased under aerobic conditions but still increased under anaerobic conditions between the second and forth week of incubation (Fig. 3 and 10). The lower amount of storage C found under aerobic conditions was accompanied by the allocation of significant proportions of C into new cell biomass. No such allocation of C to new cell biomass was found under anaerobic conditions. Given these results and the differences between the aerobic and anaerobic microbial communities discussed earlier in the text, we conclude that the aerobic microbial community has greater utilization efficiency than the anaerobic microbial community (Westermann, 1996).

	Conclusions

TOP ABSTRACT INTRODUCTION Materials and methods Results Discussion Conclusions REFERENCES

 
Nuclear magnetic resonance spectroscopy is an excellent tool for studying the microbial utilization of ¹³C–glucose in peat because it allows nondestructive assays of metabolic events. Using solution ¹³C–NMR analysis, it was possible to identify and quantify the soluble ¹³C–labeled products. Solid-state ¹³C–NMR was used to determine, quantitatively, the allocation of these carbons into new cell biomass. This overview of glucose degradation in peat would not have been possible using either one of the NMR techniques alone.

The production of storage compounds typical of fungi, such as mannitol and unsaturated triglycerides, suggested that fungi probably dominated the aerobic microbial community utilizing glucose.

The proportion of the added ¹³C used for new cell biomass (growth) under aerobic conditions was much lower than the previously reported values of between 40 and 72% for a range of soils (Bremer and Van Kessel, 1990; Bremer and Kuikman, 1994; Killham et al., 1993; Tsai et al., 1997). This indicates that peat soils may have different effects on microbial growth patterns than other minerogenic and moor humus soils.

If the ability to grow is not restricted by pH or scarcity of C, the strategy for the aerobic microbial community at high C degradation rates is to allocate most of the carbons into storage compounds. If the availability of C decreases, the storage compounds can be utilized for maintenance purposes, or for production of new cells; although, this implies that the availability of nutrients remains at a constant low level.Martin 1994; The Merck Index 1989

	ACKNOWLEDGMENTS

 
This study was supported by the Center for Environmental Research, Ume, Sweden.

Received for publication October 23, 1998.

	REFERENCES

TOP ABSTRACT INTRODUCTION Materials and methods Results Discussion Conclusions REFERENCES

 

• Baldock J.A., Oades J.M., Vassallo A.M., Wilson M.A. Incorporation of uniformly labeled ¹³C–glucose carbon into the organic fraction of a soil: Carbon balance and CP/MAS ¹³C–NMR measurements. Aust. J. Soil Res. 1989;27:725-746.
• Baldock J.A., Oades J.M., Vassallo A.M., Wilson M.A. Solid-state CP/MAS ¹³C N.M.R. analysis of bacterial and fungal cultures isolated from a soil incubated with glucose. Aust. J. Soil Res. 1990;28:213-225 a.
• Baldock J.A., Oades J.M., Vassallo A.M., Wilson M.A. Significance of microbial activity in soils as demonstrated by solid-state ¹³C NMR. Env. Sci. and Tech. 1990;24:527-530 b.
• Baldock J.A., Oades J.M., Waters A.G., Peng X., Vassallo A.M., Wilson M.A. Aspects of the chemical structure of soil organic materials as revealed by solid-state ¹³C NMR spectroscopy. Biogeochem. 1992;16:1-42.
• Baldock J.A., Preston C.M. Chemistry of carbon decomposition processes in forests as revealed by solid-state ¹³C NMR. In: Kelly J.M., McFee W.W., eds. Carbon forms and functions in forest soils. Madison, WI: SSSA, 1995:90-117 Proc. of the 8th North American Forest Soils Conf., 9–13 May 1993..
• Bengsch E., Perly B., Deleuze C., Valero A. A general rule for the assignment of the carbon-13 NMR peaks in fatty acid chains. J. Magn. Reson. 1986;68:1-13.
• Bergman I., Lundberg P., Nilsson M. Microbial carbon mineralization in an acid surface peat: Effects of changing environmental conditions for laboratory incubations. Soil Biol. Biochem. 1999;31:1867-1877.
• Bergman I., Svensson B.H., Nilsson M. Regulation of methane production in an Swedish acid mire by pH, temperature, and substrate. Soil Biol. Biochem. 1998;30:29-741.
• Bremer E., Van Kessel C. Extractability of microbial ¹⁴C and ¹⁵N following addition of variable rates of labeled glucose and (NH₄)₂SO₄ to soil. Soil Biol. Biochem 1990;22:707-713.
• Bremer E., Kuikman P. Microbial utilization of ¹⁴C[U]glucose in soil is affected by the amount and timing of glucose additions. Soil Biol. Biochem 1994;26:511-517.
• Cavigelli M.A., Robertson G.P., Klug M.J. Fatty acid methyl ester (FAME) profiles as measures of soil microbial community structure. Plant Soil 1995;170:99-113.
• Dolfing J., Tiedje J.M. Acetate inhibition of methanogenic, syntrophic benzoate degradation. Appl. Environ. Microbiol. 1988;54:1871-1873.[Abstract/Free Full Text]
• Dudley R.L., Fyfe C.A., Preston C.M. A ¹³C CPMAS NMR spectroscopic study of the transformation of plant material to peat and coal. Can. J. Spectrosc. 1990;35:31-38.
• Eurola S., Hicks S., Kaakinen E. Key to Finnish mire types. In: Moore P.D., ed. European mires. London: Academic Press, 1984:11-117.
• Fan T.W.M. Metabolic profiling by one- and two-dimensional NMR analysis of complex mixtures. Prog. Nucl. Magn. Reson. Spectrom. 1996;28:161-219.
• Golchin A., Clarke P., Oades J.M. The heterogenous nature of microbial products as shown by solid-state ¹³C CP/MAS NMR spectroscopy. Biogeochem. 1996;34:71-97.
• Gottschalk G. Bacterial fermentations. In: Mortimer P.S., ed. Bacterial metabolism. New York: Springer-Verlag, 1979:167-221.
• Harwood J.L., Russell N.J. Lipids in plants and microbes. London: Georg Allen & Unwin, 1984.
• Houwen F.P., Dijkema C., Stams A.J.M., Zehnder A.J.B. Propionate metabolism in anaerobic bacteria: Determination of carboxylation reactions with ¹³C–NMR spectroscopy. Biochim. Biophys. Acta 1991;1056:126-132.
• Holmgren A., Wikander G., Norden B. Spectroscopic investigations of peat–water interactions: An ESR, FT-IR, and NMR study. Soil Sci. 1990;149:279-291.
• Killham K., Amato M., Ladd J.N. Effect of substrate location in soil and soil pore-water regime on carbon turnover. Soil Biol. Biochem. 1993;25:57-62.
• Kotsyurbenko O.R., Nozhevnikova A.N., Soloviova T.I., Zavarzin G.A. Methanogenesis at low temperatures by microflora of tundra wetland soil. Antonie van Leeuwenhoek 1996;69:75-86.[ISI][Medline]
• Krumböck M., Conrad R. Metabolism of position-labeled glucose in anoxic methanogenic paddy soil and lake sediment. FEMS Microbiol. Ecol. 1991;85:247-256.
• Martin F., Canet D., Marchal J.P., Brondeau J. In vivo natural-abundance ¹³C nuclear magnetic resonance studies of living ectomycchorizal fungi. Plant Physiol. 1984;75:151-153.[Abstract/Free Full Text]
• Martin F., Canet D., Marchal J.P. ¹³C Nuclear magnetic resonance study of mannitol cycle and trehalose synthesis during glucose utilization by the ectomycorrhizal ascomycete cenococcum graniforme. Plant Physiol. 1985;77:499-502.[Abstract/Free Full Text]
• Martin F., Ramstedt M., Söderhäll K., Canet D. Carbohydrate and amino acid metabolism in the ectomycchorizal ascomycete Sphaerosporella brunnea during glucose utilization. Plant Physiol.. 1988;86:935-940.[Abstract/Free Full Text]
• Martin F.M. Nuclear magnetic resonance studies of ectomycorrhizal fungi. In: Norris J.R., et al. , ed. Techniques for mycorrhizal research. London: Academic Press, 1994:121-148.
• Mikkelä C., Sundh I., Svensson B.H., Nilsson M. Diurnal variation in methane emission in relation to the water table, soil temperature, climate, and vegetation cover in an Swedish acid mire. Biogeochem. 1995;28:93-114.
• Nilsson M., Bth E., Söderström B. The microfungal communities of a mixed mire in northern Sweden. Can. J. Bot. 1992;70:272-276.
• Nordén B., Bohlin E., Nilsson M., Albano ., Röckner C. Characterization of particle size fractions of peat: An integrated biological, chemical, and spectroscopic approach. Soil Sci. 1992;153:382-396.
• Preston C.M. Applications of NMR to soil organic matter analysis: History and prospects. Soil Sci. 1996;161:144-166.
• Richards B.N. The microbiology of terrestrial ecosystems. Essex, UK: Longman Group, 1987.
• Roberts, M.F., J.N.S. Evans, C.J. Tolman, and D.P. Raleigh. 1987. NMR studies of methanogens: What good is a cyclic pyrophosphate? p. 17–32. In Physiological NMR spectrocopy: From isolated cells to man. N.Y. Acad. Sci., New York.
• Rutar V., Burgar M., Blinc R. ¹³C–NMR determination of the oil composition in individual plant seeds. J. Magn. Reson. 1977;27:83-90.
• Sawyer T.E., King G.M. Glucose uptake and end product formation in an intertidal marine sediment. Appl. Environ. Microbiol. 1993;59:120-128.[Abstract/Free Full Text]
• Schink B. Principles and limitation of anaerobic degradation: Environmental and technological aspects. In: Zhender A.J.B., ed. Biology of anaerobic microorganisms. New York: Wiley, 1988:771-846.
• Schink B., Friedrich M. Energetics of syntrophic fatty acid oxidation. FEMS Microbiol. Rev. 1994;15:85-94.
• Schultz S., Conrad R. Influence of temperature on pathways to methane production in the permanently cold profundal sediment of Lake Constance. FEMS Microbiol. Ecol. 1996;20:1-14.
• Stanier R.Y., Ingraham J.L., Whedis M.L., Painter P.R. General microbiology. Englewood Cliffs, NJ: Prentice-Hall, 1986.
• Stumm W., Morgan J.J. Aquatic chemistry, 3rd ed New York: John Wiley and Sons, 1996.
• The Merck Index. 1989. p. 938. S. Budavari. (ed.) Rahway, NJ.
• Tsai C.S., Killham K., Cresser M.S. Dynamic response of microbial biomass, respiration rate, and ATP to glucose additions. Soil Biol. and Biochem. 1997;29:1249-1256.
• Vanderhart D.L., Orts W.J., Marchessault R.H. ¹³C NMR determination of the degree of cocrystallization in random copolymers of poly(ßß-hydroxyburyrate-co-ßß-hydroxyvalerate). Macromol. 1995;28:6394-6400.
• Webster E.A., Chudek J.A., Hopkins D.W. Fates of ¹³C from enriched glucose and glycine in an organic soil determined by solid-state NMR. Biol. Fertil. Soils 1997;25:389-395.
• Westermann P. The effect of incubation temperature on steady-state concentrations of hydrogen and volatile fatty acids during anaerobic degradation in slurries from wetland sediments. FEMS Microbiol. Ecol. 1994;13:295-302.
• Westermann P. Temperature regulation of anaerobic degradation of organic matter. World J. Microbiol. Biotechnol. 1996;12:497-503.
• Zelles L., Bai Q.J. Fractionation of fatty acids derived from soil lipids by solid phase extraction and their quantitative analysis by GC–MS. Soil Biol. Biochem. 1993;25:495-507.
• Zelles L., Rackwitz R., Bai Q.J., Beck T., Beese F. Discrimination of microbial diversity by fatty acid profiles of phospholipids and lipopolysaccharides in differently cultivated soils. Plant Soil 1995;170:115-122.

This Article Abstract Figures Only Full Text (PDF) Free Alert me when this article is cited Alert me if a correction is posted