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. 2009 Feb 5;75(7):1852–1859. doi: 10.1128/AEM.02745-08

In Situ Hydrogen and Nitrous Oxide as Indicators of Concomitant Fermentation and Denitrification in the Alimentary Canal of the Earthworm Lumbricus terrestris▿

Pia K Wüst 1, Marcus A Horn 1, Harold L Drake 1,*
PMCID: PMC2663201  PMID: 19201982

Abstract

The earthworm gut is a unique microzone in aerated soils that has been proposed to selectively stimulate ingested soil microorganisms by its in situ conditions, which include anoxia, high water content, a near-neutral pH, and high concentrations of organic compounds. The central objective of this study was to resolve potential links between in situ conditions and anaerobic microbial activities during the gut passage of Lumbricus terrestris. Both H2 and N2O were emitted by living earthworms, and in situ microsensor analyses revealed both H2 and N2O in the O2-free gut center. The highest H2 concentrations occurred in foregut and midgut regions, whereas the highest N2O concentrations occurred in crop/gizzard and hindgut regions. Thus, H2-producing fermentations were more localized in the foregut and midgut, whereas denitrification was more localized in the crop/gizzard and hindgut. Moisture content, total carbon, and total nitrogen were highest in the foregut and decreased from the anterior to posterior end of the gut. Nitrite, ammonium, and iron(II) concentrations were highest in the crop/gizzard and decreased from the anterior to posterior end of the alimentary canal. Concentrations of soluble organic compounds were indicative of distinct fermentation processes along the alimentary canal, with maximal concentrations of organic acids (e.g., acetate and butyrate) occurring in the midgut. These findings suggest that earthworms (i) contribute to the terrestrial cycling of carbon and nitrogen via anaerobic microbial activities in the alimentary canal and (ii) constitute a mobile source of reductant (i.e., emitted H2) for microbiota in aerated soils.

Earthworms constitute the dominant soil macrofauna in many soils (15, 34, 36) and emit the greenhouse gas nitrous oxide (N2O) concomitantly with molecular nitrogen (N2) via denitrifying bacteria in the gut (10, 11, 24, 26, 30, 39). The in situ conditions of the earthworm gut, which include anoxia, high water content, a near-neutral pH, and high concentrations of organic compounds, may activate certain ingested soil bacteria, in particular bacteria capable of anaerobiosis (10, 11, 27, 28-30, 33, 62). In this regard, the numbers of cultured anaerobes and denitrifiers are two to three orders of magnitude greater in the earthworm gut than in soil (28-30), and the amount of bacterial long-chain fatty acids is higher in the gut than in soil (46).

Gut contents of earthworms contain up to 80% intestinal mucus (i.e., up to 0.8 g mucus per g [dry weight] gut content) that consists of monosaccharides, low-molecular-weight amino acids, and glycoproteins (5, 37, 42, 62). This large amount of degradable organic carbon in the anoxic gut suggests that fermentation is very active during gut passage (27). Indeed, fermentative bacteria are abundant in the gut of earthworms (25, 28, 30). Nitrate reducers also are abundant in the earthworm gut (28) and can produce fermentation products when nitrate is not available (58). Many fermentative or facultative microorganisms also can reduce nitrite as well as nitrate (7, 63). However, fermentation processes in the alimentary canal of earthworms (Fig. 1) are not resolved, and relatively little is known about in situ conditions and microbial activities in the anterior part of the alimentary canal, i.e., in the crop and gizzard of earthworms.

FIG. 1.

FIG. 1.

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Diagram of the earthworm alimentary canal. Modified from reference 27.

Based on these collective findings, we hypothesized that fermentation is ongoing and concomitant with denitrification during gut passage. The central objective of this study was to address this hypothesis by resolving potential links between in situ conditions and anaerobic microbial activities along the alimentary canal of the anecic earthworm Lumbricus terrestris.

MATERIALS AND METHODS

Earthworms.

Adult L. terrestris L. organisms were purchased from ANZO (Bayreuth, Germany) and maintained in soil at 15°C for approximately 10 days. Soil was obtained from the meadow Trafo Wiese in Bayreuth (Germany), which is described elsewhere (27).

Emission of H2 and N2O by living earthworms.

Earthworms were washed with water, dried with tissue paper, and weighed. Two to three earthworms having a collective fresh weight of approximately 7 g were placed in a gas-tight 34-ml serum vial that then was crimp sealed under air and incubated at room temperature (approximately 20°C). The production of H2 and N2O was analyzed periodically; unless otherwise stated, values are the means from four replicates.

Microsensor measurements.

Earthworms were anesthetized with 100% carbon dioxide to prevent defecation and subsequently were sedated with 20% ethanol for approximately 15 min. Earthworms were embedded horizontally in agar (1.5%) and overlaid with agar and double-distilled water (for the measurement of N2O) or agar and 0.9% NaCl (for the measurement of H2 or O2), respectively. (Calibration tests demonstrated that these contrasting conditions were needed for microsensor stability.) This protocol was not lethal to earthworms. Microsensors for the measurement of N2O (1), H2 (67), and O2 (44) were purchased from Unisense (Aarhus, Denmark) and were mounted on a micromanipulator (Märtzhäuser, Wetzlar, Germany). Radial concentration profiles were measured at room temperature for the crop/gizzard, foregut, midgut, or hindgut region of the alimentary canal (Fig. 1). Alternatively, two-point measurements at the worm surface and the center of the alimentary canal were obtained for all four regions.

Production of H2 by dissected guts.

Earthworms were washed, sacrificed by brief immersion in 70°C water, and transferred into an O2-free chamber (Mecaplex, Grenchen, Switzerland). Guts were dissected and placed in gas-tight 34-ml serum vials (three to four gut samples with a collective fresh weight of approximately 4.5 g per vial) that were flushed with 100% argon. Guts were incubated at room temperature in the dark, and the production of H2 was analyzed periodically. Unless otherwise stated, values are means from four replicates.

Production of N2O by earthworm sections.

Earthworms were washed, dried with tissue paper, and sedated on ice. Earthworms were transferred into an O2-free chamber and separated into crop/gizzard, foregut, midgut, and hindgut sections with sterile scissors. Sections were weighed and placed in gas-tight 15-ml serum vials (two sections with a collective fresh weight of approximately 2 g per vial) that were flushed with 100% argon. Earthworm sections were incubated at room temperature in the dark. The emission of N2O was determined with and without acetylene (15%, vol/vol), an inhibitor of N2O-reductase that reduces N2O to N2 (68, 70). The emission of N2O was analyzed periodically. Unless otherwise stated, values are means from three replicates.

Extraction of soil and gut contents.

Earthworms were washed and sacrificed by brief immersion in 70°C water. A total of 15 earthworms were dissected under oxic conditions (29, 30). The alimentary canal was divided into four regions (containing crop/gizzard, foregut, midgut, or hindgut). Material from the same section of five specimens was pooled to obtain approximately 0.5 g (fresh weight) per sample. Valeric acid was added as an internal standard (32). Soil or gut content was extracted with 1.5 ml double-distilled water at 60°C by being vortexed at maximum speed for approximately 1 min. Samples subsequently were cooled on ice and homogenized with an end-over-end shaker at 5°C overnight. Solid matter was separated by centrifugation (23,700 × g, 6 min, 4°C) and used for the determination of total nitrogen, total carbon, and organic carbon contents. Supernatant fluids were filtered (0.2 μm pore size) and stored at −20°C until analyzed for soluble organic compounds, nitrate, nitrite, iron(II), and ammonium.

Analytical techniques.

The moisture content was determined by weighing the soil and gut content before and after it was dried at 60°C for 72 h. Oven-dried solid matter was ground with a ball mill (MM2; Retsch, Haan, Germany) and analyzed for total nitrogen content, and total carbon content was analyzed with an NC analyzer (Flash EA 1112; CE Instruments, Wigan, United Kingdom). Inorganic carbon was estimated as the amount of carbon lost by treatment with 8% HCl overnight followed by being at 80°C for 1 to 2 h; organic carbon was defined as the difference between total and inorganic carbon. Nitrate was analyzed with a Dionex DX-100 ion chromatograph equipped with an IonPac AS4A-SC ion-exchange column and an ED40 electrochemical detector (Sunnyvale, CA). Nitrite and iron(II) levels were determined photometrically (20, 55). Ammonium was measured by flow injection analysis (FIA-LAB; MLE, Dresden, Germany). N2O and H2 levels were determined by gas chromatography (30, 32); the rates of production were calculated by linear regression analysis. Organic acids were analyzed with a 1090 Series II high-performance liquid chromatograph (HPLC) (Hewlett-Packard Palo Alto, CA) equipped with a refractive index detector, a UV detector (210 nm) (both Series 1200; Agilent Technologies, Böblingen, Germany), and an Aminex Ion Exclusion HPX-87H column (300 by 7.8 mm; Bio-Rad, Richmond, CA). The column temperature was 60°C; the mobile phase was 4 mM H3PO4 at a flow rate of 0.8 ml min−1. Chromatograms were evaluated using the ChemStation software Rev.B.02.01 (Agilent Technologies, Böblingen, Germany). Saccharides were determined with an HPLC equipped with a Dionex ED40 electrochemical detector (100-nC range), a gold electrode, and an Ag/AgCl reference electrode (Sunnyvale, CA). The hydrolysis of poly- and oligosaccharides into monosaccharides was carried out with 4 M trifluoroacetic acid (100°C, 1 h) (49). Saccharides were separated on a CarboPac PA100 column (250 by 4 mm) and eluted with stepped NaOH gradients (Table 1) (PU-1580 intelligent HPLC pump; Jasco, Großumstadt, Germany). The column temperature was 30°C (column thermostat Jet Stream Plus; Jasco, Großumstadt, Germany). Chromatograms were monitored using the software Borwin, version 1.50 (JMBS, Grenoble, France).

TABLE 1.

Sodium hydroxide gradients used for separation of saccharides by HPLCa

Treatment Time (min) Eluent A NaOH gradient (%)b
Nonhydrolyzed 0 2
15 2
35 42
37 70
39 70
40 2
Hydrolyzed 0 3
7.5 3
18 4
26 70
29 70
30 3
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a

The flow rate was 1 ml min−1.

b

Eluent A, 0.5 M NaOH; eluent B, double-distilled H2O.

RESULTS

In vivo emission of H2 by L. terrestris and anaerobic production of H2 by dissected guts.

H2 was emitted by living earthworms at a rate that approximated 6.3 ± 4.8 nmol g (fresh weight)−1 h−1 (n = 4). Dissected guts yielded 5.9 ± 1.8 μmol H2 g (fresh weight)−1 (n = 4) after 8 h under anoxic conditions. Consistently with previous findings (30, 39), N2O also was emitted under in vivo conditions at rates that approximated 0.4 ± 0.2 nmol g (fresh weight)−1 h−1 (n = 10). These initial results suggested that the alimentary canal had a high capacity for fermentative activity, and that this activity manifested itself in the in vivo emission of H2 concomitantly with the emission of N2O.

In situ concentrations of H2, N2O, and O2.

In situ H2 and N2O concentrations were highest in the O2-free core of crop/gizzard radial profiles (Fig. 2). O2 was not detected in the crop/gizzard, foregut, midgut, or hindgut (Fig. 2 and data not shown). N2O measurements were initiated at the cuticle, since preliminary tests demonstrated that the levels of N2O above the cuticle were below the detection limit. The steep decline of N2O toward the cuticle, which has been documented previously (27), suggests that the cuticle is a diffusion barrier.

FIG. 2.

FIG. 2.

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Radial microsensor profiles of O2, N2O, and H2 for the crop/gizzard of sedated L. terrestris worms. The worm radius equals 100% (as defined in the figure). Representative patterns of replicate analyses are shown.

Maximal concentrations of H2 occurred in the center of the foregut (5 μM) and midgut (6 μM) (Fig. 3). Despite the high variability among replicates, (i) the pattern of gases in a given worm section was similar, and (ii) H2, an indicator of ongoing fermentation, was detected in every replicate. Maximal concentrations of N2O occurred in the center of the crop/gizzard (2 μM) and hindgut (1 μM) (Fig. 3). The average N2O concentration at the worm surface of the midgut region was higher than that in the worm center.

FIG. 3.

FIG. 3.

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In situ concentrations of N2O and H2 along the alimentary canal of L. terrestris. Values are the means from 6 to 23 replicates; error bars indicate positive standard deviations.

Denitrification capacities of earthworm sections.

All earthworm sections emitted N2O without appreciable delay under anoxic conditions (Fig. 4 and data not shown). Crop/gizzard and hindgut sections emitted the largest amounts of N2O (Fig. 4), which is consistent with the results obtained by N2O microsensor analysis (Fig. 3). N2O emission rates increased by factors of approximately 2, 3, 4, and 5 for crop/gizzard, foregut, midgut, and hindgut, respectively, when earthworm sections were exposed to acetylene (Fig. 4), indicating that (i) N2O was derived from denitrification, (ii) N2 production exceeded N2O production, and (iii) the relative amount of N2 increased from the anterior to the posterior end of the alimentary canal.

FIG. 4.

FIG. 4.

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Production of N2O by earthworm sections with and without acetylene. Values are the means from triplicates; error bars indicate positive standard deviations.

In situ parameters potentially linked to anaerobic activities along the alimentary canal.

The results described above indicated that distinct anaerobic microbial processes occurred at different places along the alimentary canal. In situ parameters were evaluated to further resolve these microbial processes.

Moisture content along the alimentary canal was approximately twofold greater than that of the soil in which the worms were incubated (Table 2). Moisture content, total nitrogen, and total carbon content were highest in the foregut and decreased from the anterior to the posterior end of the gut. Compared to the amounts in the soil, the amounts of total nitrogen, total carbon, and organic carbon were enriched in all portions of the alimentary canal. Approximately 90% of the total carbon was organic in crop/gizzard content, a value somewhat lower than those of other earthworm regions and soil. Excreted calcium carbonate in the esophagus followed by the release of mucus into the foregut (15) may have accounted for the higher relative amount of inorganic carbon in crop/gizzard contents. The concentrations of nitrite were greater in the aqueous phase of crop/gizzard contents and foregut contents than in the aqueous phase of soil. In contrast, nitrate concentrations were highest in soil, indicating that nitrate-reducing organisms were active in the crop/gizzard and other regions of the alimentary canal. The concentrations of iron(II) and ammonium were highest in crop/gizzard content and decreased along the alimentary canal. The concentration of iron(II) at the end of gut passage was equivalent to that of soil.

TABLE 2.

In situ parameters along the alimentary canal of L. terrestrisa

Alimentary canal site Moisture content (%) Total nitrogen (mg g [dry wt]−1) Total carbon (mg g [dry wt]−1) Organic carbon (mg g [dry wt]−1) Nitrate (mmol liter [water content]−1) Nitrite (mmol liter [water content]−1) Ammonium (mmol liter [water content]−1) Iron(II) (mmol liter [water content]−1)
Crop/gizzard 41.8 ± 3.0 6.8 ± 0.5 54.9 ± 5.1 49.5b 0.3 ± 0.1 3.8 ± 0.2 17.9c 5.4 ± 0.5
Foregut 51.9 ± 0.2 9.0 ± 0.4 62.0 ± 3.5 58.8 ± 2.8 0.1 ± 0.0 2.5 ± 0.5 15.8b 2.9 ± 0.5
Midgut 49.1 ± 2.3 5.2 ± 1.2 49.6 ± 8.5 48.4 ± 8.2 0.1 ± 0.0 0.2 ± 0.1 11.0 ± 1.6 0.5 ± 0.3
Hindgut 44.0 ± 3.3 4.4 ± 0.8 42.1 ± 6.5 41.3 ± 5.9 0.3 ± 0.0 0 ± 0 11.5 ± 1.4 0.1 ± 0.1
Soil 19.9 ± 1.4 3.4 ± 0.8 38.6 ± 8.6 37.3 ± 8.2 0.9 ± 0.1 0.2 ± 0.1 0.8 ± 0.2 0.1c
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a

Unless otherwise stated, values are the means from triplicates (± standard deviations); each worm replicate consists of gut content derived from five worms.

b

No standard deviation is given because three replicates had to be pooled to obtain sufficient material for analysis.

c

No standard deviation is given because two replicates were analyzed, and the value is the average for the replicates.

The alimentary canal contained high concentrations of soluble organic compounds. Maltose, isomaltose, and glucose were dominant nonpolymeric saccharides in the aqueous phase of the alimentary canal (Fig. 5). The numbers and concentrations of detected monosaccharides increased significantly after hydrolysis with trifluoroacetic acid, indicating that saccharides in the earthworm gut were primarily polymeric. After hydrolysis, approximately 110 mM monosaccharide equivalents were obtained from crop/gizzard contents, with mannose, glucose, galactose, arabinose, rhamnose, and fucose being dominant monosaccharides. Concentrations of saccharides decreased from the anterior to the posterior end of the alimentary canal, indicating that saccharides were consumed during gut passage.

FIG. 5.

FIG. 5.

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Concentrations of saccharides along the alimentary canal of L. terrestris and in soil. I, not hydrolyzed; II, after hydrolysis with trifluoroacetic acid. Values are the means of triplicates; each replicate consists of gut content derived from five worms.

The highest concentrations (approximately 32 mM total) and highest diversity of organic acids were detected in the aqueous phase of midgut content, indicating that fermentative organisms were very active in this region of the alimentary canal (Fig. 6). Lactate and propionate occurred primarily in crop/gizzard contents and foregut contents, whereas butyrate and acetate dominated in mid- and hindgut contents. Methylbutyrate was likewise a major product in the midgut.

FIG. 6.

FIG. 6.

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Concentrations of fatty acids along the alimentary canal of L. terrestris and in soil. Values are the means from triplicates; each replicate consists of gut content derived from five worms.

DISCUSSION

Impact of in situ parameters on the microbiota along the alimentary canal of L. terrestris.

Cultured numbers of microbes capable of growth under anoxic conditions are higher in the earthworm gut than in soil (28-30), and it has been proposed that the in situ conditions of the earthworm gut activate ingested soil bacteria (11, 27, 33). The pH varies relatively little along the alimentary canal of L. terrestris (22). Intestinal mucus concentrations of earthworm species decrease along the alimentary canal (5, 62), suggesting that in situ conditions change during gut passage. The moisture content, total nitrogen, and total carbon of gut contents were greater than those of surrounding soil and decreased from the anterior to the posterior end of the gut (Table 2), findings that are consistent with previous studies (5, 27). The levels of iron(II), ammonium, and nitrite decreased along the alimentary canal (Table 2), underscoring the likelihood that ingested microorganisms are subjected to changing in situ conditions during gut passage. Free O2 was not detected in the crop/gizzard (Fig. 2), suggesting that anaerobic activities are favored immediately after soil enters the alimentary canal. The absence of free O2 in the alimentary canal and worm tissues suggests that it was rapidly bound by hemoglobin (15) and subject to respiration.

Total levels of organic carbon are higher in the earthworm alimentary canal than in soil (5, 27) (Table 2). The hydrolysis of the aqueous phase of alimentary canal contents yielded mannose, glucose, galactose, arabinose, rhamnose, and fucose (Fig. 5). The hydrolysis of the plant heteropolymer hemicellulose typically yields the pentoses xylose and arabinose, the hexoses glucose, mannose, and galactose, and the disaccharide cellobiose (45), and fucose occurs in plant cell walls, bacterial extracellular polysaccharides, and animal glycoproteins (66). Thus, at least some of the monosaccharides detected in the earthworm gut might be derived from ingested biomass. However, the earthworm gut contains up to 80% intestinal mucus (37, 42, 62), and it is likely that most of the monosaccharide equivalents detected were mucus derived. Indeed, glucosamine, galactosamine, glucose, galactose, mannose, and fucose are components of mucopolysaccharides from Lumbricus spp. (42). Polysaccharide-degrading enzymes (e.g., amylase, cellulase, xylanase, and chitinase) occur in the earthworm gut (40, 59, 61, 64, 65). The highest amylase and cellulase activities in the anterior part of the alimentary canal (59, 61) coincide with the highest organic carbon and saccharide concentrations in the crop/gizzard and foregut (Table 2 and Fig. 5). Monosaccharide-releasing enzymes (e.g., mannosidase, galactosidase, and glucosidase) are active in the earthworm gut (35) and might contribute to the occurrence of monosaccharides in the alimentary canal. These collective findings reinforce the hypothesis that the in situ conditions in the earthworm alimentary canal are ideally suited for heterotrophs capable of anaerobiosis.

Concomitant fermentation and denitrification in the alimentary canal.

H2 was emitted in vivo and produced in situ (Fig. 2 and 3). The decomposition of organic matter via fermentation can result in the formation of H2 (53). That the decrease in the amount of saccharides along the alimentary canal (Fig. 5) was concomitant to the production of H2 and other fermentation products (Fig. 3 and 6) supports the conclusion that fermentation is ongoing in the earthworm gut. Fungal hyphae are either disrupted mechanically in the anterior part of the alimentary canal or digested during gut passage (50, 51), suggesting that bacteria play a dominant role in gut fermentation (27, 28). Propionate was detected primarily in soil, crop/gizzard, and foregut (Fig. 6). Propionate is a fermentation product of propionibacteria that belong to the actinobacteria (9), and 16S rRNA gene sequences indicative of actinobacteria have been detected in casts and in the gut of Lumbricus rubellus (54). Propionibacteria ferment monosaccharides but also can utilize lactate, which was detected in the crop/gizzard and foregut. Lactic acid bacteria form a large fraction of the cultivable gut microbiota of wood- and soil-feeding termites (2, 52, 57). Most lactic acid bacteria are aerotolerant (8, 56) and, thus, might be poised to react quickly to anoxia after ingestion. Indeed, the detection of lactate primarily in the crop/gizzard (Fig. 6) suggests that it is formed in the anterior part of the alimentary canal and is subject to consumption in subsequent regions of the gut, which is consistent with the high capacity of gut homogenates to produce lactate and subsequently consume it under anoxic conditions (29).

High concentrations of H2 occurred concomitantly with formate, acetate, and succinate in the foregut and midgut. Formate, acetate, succinate, and H2 are characteristic products of facultative aerobes, including Enterobacteriaceae that are capable of fermenting various monosaccharides (e.g., glucose, fucose, and rhamnose (4, 19). Enterobacteriaceae-related species and other Gammaproteobacteria have been isolated from the gut of Aporrectodea caliginosa (28), and 16S rRNA gene sequences indicative of Gammaproteobacteria were retrieved from the gut of L. rubellus (54). The H2-producing fermenter Paenibacillus terrae MH72 (of the Firmicutes) was isolated from A. caliginosa (25). Methylbutyrate was detected in the midgut. Methylbutyrate often is produced when amino acids (e.g., valine and isoleucine) are fermented (17), suggesting that amino acids that are abundant in the earthworm gut (11, 27) were utilized as carbon sources. Products indicative of butyrate fermentation (i.e., butyrate and H2) were detected in the midgut and hindgut. Butyrate was produced in anoxic most-probable-number dilutions of gut homogenates obtained from A. caliginosa (28). Clostridiaceae-related species have been isolated from the gut of the earthworm A. caliginosa (28). Hydrogenases of butyrate-producing clostridia are O2 sensitive (6), suggesting that this functional group is not active immediately upon ingestion. Thus, a broad diversity of bacteria might be linked to the anaerobic degradation of organic matter in the earthworm gut.

The proposal that denitrification and the dissimilatory reduction of nitrate are active in the earthworm gut (27, 28) is supported by (i) the near absence of nitrate in the alimentary canal (Table 2), (ii) the increased concentration of nitrite in crop/gizzard and foregut contents compared to that of soil (Table 2), (iii) the occurrence of N2O in all regions of the anoxic alimentary canal (Fig. 3), and (iv) the acetylene-dependent enhancement of N2O production by earthworm sections (Fig. 4). In situ concentrations of N2O (Fig. 3) and N2O production by worm sections (Fig. 4) suggest that denitrification is more localized in the crop/gizzard and hindgut. Complete denitrification (i.e., the reduction of nitrate or nitrite to N2) in the crop/gizzard indicates that denitrifiers were activated without significant delay upon ingestion. The molecular analysis of nosZ (a structural gene for N2O reductase [70]) in earthworm gut contents indicated that species related to Bradyrhizobium, Flavobacterium, Dechloromonas, Brucella, Sinorhizobium, Pseudomonas, Ralstonia, and Paracoccus are involved in gut denitrification (24).

Gaseous emission in the context of worm respiration.

The respiration rate of L. terrestris approximates 3 μmol O2 g (fresh weight)−1 h−1 (34), suggesting that 12 μmol reducing equivalents g (fresh weight)−1 h−1 is directed toward the reduction of O2 by the worm. H2 was emitted by living earthworms at a rate that approximated 6 nmol g (fresh weight)−1 h−1, a value corresponding to 12 nmol reducing equivalents g (fresh weight)−1 h−1. N2O was emitted at rates that approximated 0.4 nmol g (fresh weight)−1 h−1. Assuming that the amount of N2 emitted was equal to that of N2O (26), approximately 7 nmol reducing equivalents g (fresh weight)−1 h−1 was directed toward the emission of nitrogenous gases. These values indicate that the amount of reductant lost as emitted H2 and nitrogenous gases is minimal compared to the overall flow of reductant toward the respiration of the earthworm. The millimolar amounts of organic compounds in the alimentary canal support this conclusion, i.e., that the main flow of reductant at the level of the system is not toward denitrification and H2 production. It is obvious that a serious loss of reductant (i.e., source of energy) would be problematic for the earthworm.

Trophic links along the alimentary canal.

A hypothetical model of anaerobic processes and potential trophic links along the alimentary canal of L. terrestris is proposed (Fig. 7). L. terrestris is anecic (i.e., lives in deeper soil zones, ingests moderate amounts of mineral soil, and feeds on litter dragged into its burrow) and was selected as a model earthworm in this study. However, the feeding habits of endogeic (feed in the rhizosphere, ingest substantial amounts of mineral soil, and preferentially live in upper mineral soil; e.g., A. caliginosa) and epigeic (feed on litter and preferentially live above the mineral soil; e.g., L. rubellus) earthworms may result in different processes along the alimentary canal than those shown in the model. Nonetheless, the model serves to illustrate the spatial differences among anaerobic microbial processes that might occur during gut passage.

FIG. 7.

FIG. 7.

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Hypothetical model of links between in situ conditions and anaerobic activities during the gut passage of L. terrestris. Concentrations are indicated by the font size. The tapering off of a shaded element indicates that the item identified decreases in quantity in the direction of the taper. Gases in clouds indicate the in vivo emission by the worm.

The highest in situ H2 concentrations occurred in foregut and midgut regions. In contrast, the highest in situ N2O concentrations occurred in crop/gizzard and hindgut regions (crop/gizzard and hindgut sections likewise produced the largest amount of N2O under anoxic conditions). Thus, H2-producing fermentations were localized primarily in the foregut and midgut, whereas denitrification was localized primarily in the crop/gizzard and hindgut. Oxidizable carbon is essential to both denitrifiers and fermenters. Total organic carbon and saccharides decrease along the alimentary canal, indicating that organic compounds are utilized by microorganisms during gut passage. However, organic compounds likely are subject to assimilation by the earthworm, since (i) long-chain fatty acids derived from ingested bacteria can be assimilated into earthworm tissue (46), and (ii) the capacity to assimilate organic molecules in the gastrointestinal tract is a general trait of animals (3, 13, 16, 23, 43, 60). Thus, it is very likely that fermenters, denitrifiers, and the earthworm compete for available carbon. Indeed, the capacity of the earthworm to assimilate organic compounds derived from the breakdown of intestinal mucus and ingested biomass is essential to its survival (15, 48). The occurrence of large amounts of iron(II) in the crop/gizzard suggests that iron(III) reduction occurred during the initial ingestion phase; the decreasing amount of iron(II) in the posterior regions of the gut suggests that it is subject to assimilation by either microorganisms or the worm during gut passage.

Numerous anaerobes occur in the earthworm gut (29, 30) and are proposed to be the main source of organic acids in the alimentary canal (Fig. 6). Fermentation products indicative of distinct fermentations occurred spatially and consequently in temporal sequence in the alimentary canal. Total cell counts and numbers of cultured bacteria increase from foregut to hindgut (18, 31, 51). Long-chain fatty acid profiles and cell numbers for bacterial phyla (i.e., Alpha-, Beta-, Gamma-, and Deltaproteobacteria) detected by fluorescent in situ hybridization differ significantly along the earthworm alimentary canal of L. terrestris (47, 50). These collective findings indicate that both the microbial community structure and associated activities change during gut passage.

H2 was a stable end product in glucose-supplemented anoxic gut homogenates (29) and was produced by gut sections and living earthworms. Methane is neither emitted by earthworms nor formed by gut homogenates (29). Thus, although H2 forms important trophic links to methanogenesis and acetogenesis (12, 21, 69), methanogens and acetogens are not metabolically significant in the earthworm gut (29). In contrast, the specialized digestive system of termites yields large amounts of H2 that are tightly linked to acetogenesis in the hindgut paunch (41). Although it is currently unknown if there are reductant sinks for H2 in the earthworm alimentary canal, the occurrence of different inorganic electron acceptors (Table 2) suggests that it is subject to consumption as well as emission.

Living earthworms (i.e., L. terrestris) emitted approximately 6 nmol H2 g (fresh weight)−1 h−1. Assuming worm densities of up to 2,000 individuals per square meter (14, 15) and an average worm weight of 2 g, earthworms can emit up to 600 μmol H2 per square meter per day. This value for H2 is sevenfold higher than the daily emission of N2O per square meter of pasture (38). Thus, earthworms might constitute a mobile source of reductant (i.e., emitted H2) for the microbiota in aerated soils.

Acknowledgments

We thank Tanja Gonter and Christine Stöcker (Central Analytic Laboratory, University of Bayreuth) for excellent technical assistance and Matthias Lorenz (Department of Animal Ecology, University of Bayreuth) for providing the HPLC and advice on saccharide analysis.

Support for this study was provided by grants from the Deutsche Forschungsgemeinschaft (DR310/4-1) and the University of Bayreuth.

Footnotes

▿

Published ahead of print on 5 February 2009.

REFERENCES

  • 1.Andersen, K., T. Kjaer, and N. P. Revsbech. 2001. An oxygen insensitive microsensor for nitrous oxide. Sens. Actuators B Chem. 81:42-48. [Google Scholar]
  • 2.Bauer, S., A. Tholen, J. Overmann, and A. Brune. 2000. Characterization of abundance and diversity of lactic acid bacteria in the hindgut of wood- and soil-feeding termites by molecular and culture-dependent techniques. Arch. Microbiol. 173:126-137. [DOI] [PubMed] [Google Scholar]
  • 3.Bergman, E. N. 1990. Energy contributions of volatile fatty-acids from the gastrointestinal-tract in various species. Physiol. Rev. 70:567-590. [DOI] [PubMed] [Google Scholar]
  • 4.Brenner, D. J. 1992. Introduction to the family Enterobacteriaceae, p. 2673-2695. In A. Balows, H. G. Trüper, M. Dworkin, W. Harder, and K.-H. Schleifer (ed.), The prokaryotes, 2nd ed. Springer Verlag, New York, NY.
  • 5.Brown, G. G., I. Barois, and P. Lavelle. 2000. Regulation of soil organic matter dynamics and microbial activity in the drilosphere and the role of interactions with other edaphic functional domains. Eur. J. Soil Biol. 36:177-198. [Google Scholar]
  • 6.Buckel, W. 2005. Special clostridial enzymes and fermentation pathways, p. 122-220. In P. Dürre (ed.), Handbook on clostridia. CRC Press, Boca Raton, FL.
  • 7.Cole, J. A. 1990. Physiology, biochemistry, and genetics of nitrate dissimilation to ammonia, p. 57-76. In N. P. Revsbech and J. Sørensen (ed.), Denitrification in soils and sediment. Plenum Press, New York, NY.
  • 8.Condon, S. 1987. Responses of lactic-acid bacteria to oxygen. FEMS Microbiol. Rev. 46:269-280. [Google Scholar]
  • 9.Cummins, C. S., and J. L. Johnson. 1992. The genus Propionibacterium, p. 834-849. In A. Balows, H. G. Trüper, M. Dworkin, W. Harder, and K.-H. Schleifer (ed.), The prokaryotes, 2nd ed. Springer, New York, NY.
  • 10.Drake, H. L., and M. A. Horn. 2006. Earthworms as a transient heaven for terrestrial denitrifying microbes: a review. Eng. Life Sci. 6:261-265. [Google Scholar]
  • 11.Drake, H. L., and M. A. Horn. 2007. As the worm turns: the earthworm gut as a transient habitat for soil microbial biomes. Annu. Rev. Microbiol. 61:169-189. [DOI] [PubMed] [Google Scholar]
  • 12.Drake, H. L., K. Küsel, and C. Matthies. 2006. Acetogenic prokaryotes, p. 354-420. In A. Balows, H. G. Trüper, M. Dworkin, W. Harder, and K.-H. Schleifer (ed.), The prokaryotes, 3rd ed. Springer, New York, NY.
  • 13.Duke, G. E. 1997. Gastrointestinal physiology and nutrition in wild birds. Proc. Nutr. Soc. 56:1049-1056. [DOI] [PubMed] [Google Scholar]
  • 14.Edwards, C. A. 2004. The importance of earthworms as key representatives of the soil fauna, p. 3-11. In C. A. Edwards (ed.), Earthworm ecology, 2nd ed. CRC Press, Boca Raton, FL.
  • 15.Edwards, C. A., and P. J. Bohlen. 1996. Biology and ecology of earthworms, 3rd ed. Chapman & Hall, London, United Kingdom.
  • 16.Engelhardt, W. V., and G. Rechkemmer. 1985. Colonic transport of short-chain fatty acids and the importance of microclimate, p. 93-101. In E. Skadhauge and K. Heintz (ed.), Intestinal absorption and secretion. MTP Press, Lancaster, United Kingdom.
  • 17.Fardeau, M. L., B. K. Patel, M. Magot, and B. Ollivier. 1997. Utilization of serine, leucine, isoleucine, and valine by Thermoanaerobacter brockii in the presence of thiosulfate or Methanobacterium sp. as electron acceptors. Anaerobe 3:405-410. [DOI] [PubMed] [Google Scholar]
  • 18.Fischer, K., D. Hahn, R. I. Amann, O. Daniel, and J. Zeyer. 1995. In situ analysis of the bacterial community in the gut of the earthworm Lumbricus terrestris L by whole-cell hybridization. Can. J. Microbiol. 41:666-673. [Google Scholar]
  • 19.Gottschalk, G. 1986. Bacterial metabolism, 2nd ed. Springer, New York, NY.
  • 20.Harrigan, W. F., and M. E. McCance. 1966. Laboratory methods in microbiology. Academic Press, London, United Kingdom.
  • 21.Hedderich, R., and W. B. Whitman. 2006. Physiology and biochemistry of the methane-producing Archaea, p. 1050-1079. In M. M. Dworkin, S. Falkow, E. Rosenberg, K.-H. Schleifer, and E. Stackebrandt (ed.), The prokaryotes, 3rd ed., vol. 2. Springer, New York, NY. [Google Scholar]
  • 22.Heran, H. 1956. Ein Beitrag zur Verdauungsphsiologie von Lumbricus terrestris L. J. Comp. Physiol. A 39:44-62. [Google Scholar]
  • 23.Hogan, M. E., M. Slaytor, and R. W. Obrien. 1985. Transport of volatile fatty-acids across the hindgut of the cockroach Panesthia cribrata Saussure and the termite Mastotermes darwiniensis Froggatt. J. Insect Physiol. 31:587-591. [Google Scholar]
  • 24.Horn, M. A., H. L. Drake, and A. Schramm. 2006. Nitrous oxide reductase genes (nosZ) of denitrifying microbial populations in soil and the earthworm gut are phylogenetically similar. Appl. Environ. Microbiol. 72:1019-1026. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Horn, M. A., J. Ihssen, C. Matthies, A. Schramm, G. Acker, and H. L. Drake. 2005. Dechloromonas denitrificans sp. nov., Flavobacterium denitrificans sp. nov., Paenibacillus anaericanus sp. nov. and Paenibacillus terrae strain MH72, N2O-producing bacteria isolated from the gut of the earthworm Aporrectodea caliginosa. Int. J. Syst. Evol. Microbiol. 55:1255-1265. [DOI] [PubMed] [Google Scholar]
  • 26.Horn, M. A., R. Mertel, M. Kästner, M. Gehre, and H. L. Drake. 2006. In vivo emission of dinitrogen by earthworms via denitrifying bacteria in the gut. Appl. Environ. Microbiol. 72:1013-1018. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Horn, M. A., A. Schramm, and H. L. Drake. 2003. The earthworm gut: an ideal habitat for ingested N2O-producing microorganisms. Appl. Environ. Microbiol. 69:1662-1669. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 28.Ihssen, J., M. A. Horn, C. Matthies, A. Gößner, A. Schramm, and H. L. Drake. 2003. N2O-producing microorganisms in the gut of the earthworm Aporrectodea caliginosa are indicative of ingested soil bacteria. Appl. Environ. Microbiol. 69:1655-1661. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Karsten, G. R., and H. L. Drake. 1995. Comparative assessment of the aerobic and anaerobic microfloras of earthworm guts and forest soils. Appl. Environ. Microbiol. 61:1039-1044. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 30.Karsten, G. R., and H. L. Drake. 1997. Denitrifying bacteria in the earthworm gastrointestinal tract and in vivo emission of nitrous oxide (N2O) by earthworms. Appl. Environ. Microbiol. 63:1878-1882. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Krištůfek, V., K. Ravasz, and V. Pižl. 1992. Changes in densities of bacteria and microfungi during gut transit in Lumbricus rubellus and Aporrectodea caliginosa (Oligochaeta, Lumbricidae). Soil Biol. Biochem. 24:1499-1500. [Google Scholar]
  • 32.Küsel, K., and H. L. Drake. 1995. Effects of environmental parameters on the formation and turnover of acetate by forest soils. Appl. Environ. Microbiol. 61:3667-3675. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Lavelle, P., C. Lattaud, D. Trigo, and I. Barois. 1995. Mutualism and biodiversity in soils. Plant Soil. 170:23-33. [Google Scholar]
  • 34.Lee, K. E. 1985. Earthworms. Academic Press, Sydney, Australia.
  • 35.Li, Y. T., and M. R. Shetlar. 1965. Glycosidases in earthworm Lumbricus terrestris. Comp. Biochem. Physiol. 14:275-279. [DOI] [PubMed] [Google Scholar]
  • 36.Makeschin, F. 1997. Earthworms (Lumbricidae: Oligochaeta): important promoters of soil development and soil fertility, p. 173-223. In G. Benckiser (ed.), Fauna in soil ecosystems. Marcel Dekker Inc., New York, NY.
  • 37.Martin, A., J. Cortez, I. Barois, and P. Lavelle. 1987. The production of intestinal mucus by earthworms—a key process in their interactions with the soil microflora. Rev. Ecol. Biol. Sol. 24:549-558. [Google Scholar]
  • 38.Matson, P. A., P. M. Vitousek, G. P. Livingston, and N. A. Swanberg. 1990. Sources of variation in nitrous oxide flux from Amazonian ecosystems. J. Geophys. Res. Atmos. 95:16789-16798. [Google Scholar]
  • 39.Matthies, C., A. Griesshammer, M. Schmittroth, and H. L. Drake. 1999. Evidence for involvement of gut-associated denitrifying bacteria in emission of nitrous oxide (N2O) by earthworms obtained from garden and forest soils. Appl. Environ. Microbiol. 65:3599-3604. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40.Mishra, P. C., and M. C. Dash. 1980. Digestive enzymes of some earthworms. Experientia 36:1156-1157. [DOI] [PubMed] [Google Scholar]
  • 41.Pester, M., and A. Brune. 2007. Hydrogen is the central free intermediate during lignocellulose degradation by termite gut symbiosis. ISME J. 1:551-565. [DOI] [PubMed] [Google Scholar]
  • 42.Rahemtulla, F., and S. Lovtrup. 1975. Comparative biochemistry of invertebrate mucopolysaccharides. 3. Oligochaeta and Hirudinea. Comp. Biochem. Physiol. B Biochem. Mol. Biol. 50:627-629. [DOI] [PubMed] [Google Scholar]
  • 43.Rechkemmer, G., K. Ronnau, and W. von Engelhardt. 1988. Fermentation of polysaccharides and absorption of short chain fatty acids in the mammalian hindgut. Comp. Biochem. Physiol. A Comp. Physiol. 90:563-568. [DOI] [PubMed] [Google Scholar]
  • 44.Revsbech, N. P. 1989. An oxygen microsensor with a guard cathode. Limnol. Oceanogr. 34:474-478. [Google Scholar]
  • 45.Saddler, J. N., E. K. C. Yu, M. Meshartree, N. Levitin, and H. H. Brownell. 1983. Utilization of enzymatically hydrolyzed wood hemicelluloses by microorganisms for production of liquid fuels. Appl. Environ. Microbiol. 45:153-160. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46.Sampedro, L., R. Jeannotte, and J. K. Whalen. 2006. Trophic transfer of fatty acids from gut microbiota to the earthworm Lumbricus terrestris L. Soil Biol. Biochem. 38:2188-2198. [Google Scholar]
  • 47.Sampedro, L., and J. K. Whalen. 2007. Changes in the fatty acid profiles through the digestive tract of the earthworm Lumbricus terrestris L. Appl. Soil Ecol. 35:226-236. [Google Scholar]
  • 48.Scheu, S. 1991. Mucus excretion and carbon turnover of endogenic earthworms. Biol. Fertil. Soils 12:217-220. [Google Scholar]
  • 49.Schiller, M., H. von der Heydt, F. Marz, and P. C. Schmidt. 2002. Quantification of sugars and organic acids in hygroscopic pharmaceutical herbal dry extracts. J. Chromatogr. A 968:101-111. [DOI] [PubMed] [Google Scholar]
  • 50.Schönholzer, F., D. Hahn, B. Zarda, and J. Zeyer. 2002. Automated image analysis and in situ hybridization as tools to study bacterial populations in food resources, gut and cast of Lumbricus terrestris L. J. Microbiol. Methods 48:53-68. [DOI] [PubMed] [Google Scholar]
  • 51.Schönholzer, F., D. Hahn, and J. Zeyer. 1999. Origins and fate of fungi and bacteria in the gut of Lumbricus terrestris L. studied by image analysis. FEMS Microbiol. Ecol. 28:235-248. [Google Scholar]
  • 52.Schultz, J. E., and J. A. Breznak. 1978. Heterotrophic bacteria present in hindguts of wood-eating termites [Reticulitermes flavipes (Kollar)]. Appl. Environ. Microbiol. 35:930-936. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 53.Schwartz, E., and B. Friedrich. 2006. The H2-metabolizing prokaryotes, p. 496-563. In A. Balows, H. G. Trüper, M. Dworkin, W. Harder, and K.-H. Schleifer (ed.), The prokaryotes, 3rd ed. Springer, New York, NY.
  • 54.Singleton, D. R., P. F. Hendrix, D. C. Coleman, and W. B. Whitman. 2003. Identification of uncultured bacteria tightly associated with the intestine of the earthworm Lumbricus rubellus (Lumbricidae; Oligochaeta). Soil Biol. Biochem. 35:1547-1555. [Google Scholar]
  • 55.Tamura, H., K. Goto, T. Yotsuyan, and M. Nagayama. 1974. Spectrophotometric determination of iron(II) with 1,10-phenanthroline in presence of large amounts of iron(III). Talanta 21:314-318. [DOI] [PubMed] [Google Scholar]
  • 56.Teuber, M., A. Geis, and H. Neve. 1992. The genus Lactococcus, p. 1482-1501. In A. Balows, H. G. Trüper, M. M. Dworkin, W. Harder, and K.-H. Schleifer (ed.), The prokaryotes, 2nd ed., vol. 2. Springer, New York, NY. [Google Scholar]
  • 57.Tholen, A., B. Schink, and A. Brune. 1997. The gut microflora of Reticulitermes flavipes, its relation to oxygen, and evidence for oxygen-dependent acetogenesis by the most abundant Enterococcus sp. FEMS Microbiol. Ecol. 24:137-149. [Google Scholar]
  • 58.Tiedje, J. M. 1988. Ecology of denitrification and dissimilatory nitrate reduction to ammonium, p. 179-243. In A. J. B. Zehnder (ed.), Biology of anaerobic microorganisms. John Wiley and Sons, New York, NY.
  • 59.Tillinghast, E. K., R. O'Donnell, D. Eves, E. Calvert, and J. Taylor. 2001. Water-soluble luminal contents of the gut of the earthworm Lumbricus terrestris L. and their physiological significance. Comp. Biochem. Physiol. A Comp. Physiol. 129:345-353. [DOI] [PubMed] [Google Scholar]
  • 60.Titus, E., and G. A. Ahearn. 1991. Transintestinal acetate transport in a herbivorous teleost: anion-exchange at the basolateral membrane. J. Exp. Biol. 156:41-61. [Google Scholar]
  • 61.Tracey, M. V. 1951. Cellulase and chitinase of earthworms. Nature 167:776-777. [DOI] [PubMed] [Google Scholar]
  • 62.Trigo, D., I. Barois, M. H. Garvin, E. Huerta, S. Irisson, and P. Lavelle. 1999. Mutualism between earthworms and soil microflora. Pedobiologia 43:866-873. [Google Scholar]
  • 63.Umarov, M. M. 1990. Biotic sources of nitrous oxide in the context of the global budget of nitrous oxide, p. 263-268. In A. F. Bouwman (ed.), Soils and the greenhouse effect. John Wiley and Sons, Chichester, United Kingdom.
  • 64.Urbášek, F. 1990. Cellulase activity in the gut of some earthworms. Rev. Ecol. Biol. Sol. 27:21-28. [Google Scholar]
  • 65.Urbášek, F., and V. Pižl. 1991. Activity of digestive enzymes in the gut of 5 earthworm species (Oligochaeta, Lumbricidae). Rev. Ecol. Biol. Sol. 28:461-468. [Google Scholar]
  • 66.Vanhooren, P. D., and E. J. Vandamme. 1999. L-fucose: occurrence, physiological role, chemical, enzymatic and microbial synthesis. J. Chem. Technol. Biotechnol. 74:479-497. [Google Scholar]
  • 67.Witty, J. F. 1991. Microelectrode measurements of hydrogen concentrations and gradients in legume nodules. J. Exp. Bot. 42:765-771. [Google Scholar]
  • 68.Yoshinari, T., and R. Knowles. 1976. Acetylene inhibition of nitrous oxide reduction by denitrifying bacteria. Biochem. Biophys. Res. Commun. 69:705-710. [DOI] [PubMed] [Google Scholar]
  • 69.Zinder, S. H. 1993. Physiological ecology of methanogens, p. 128-206. In J. G. Ferry (ed.), Methanogenesis. Chapman & Hall, New York, NY.
  • 70.Zumft, W. G. 1997. Cell biology and molecular basis of denitrification. Microbiol. Mol. Biol. Rev. 61:533-616. [DOI] [PMC free article] [PubMed] [Google Scholar]

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