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Host-Microbial Interactions
Research Article
13 July 2021

Methanogenesis in the Digestive Tracts of the Tropical Millipedes Archispirostreptus gigas (Diplopoda, Spirostreptidae) and Epibolus pulchripes (Diplopoda, Pachybolidae)


Methanogens represent the final decomposition step in anaerobic degradation of organic matter, occurring in the digestive tracts of various invertebrates. However, factors determining their community structure and activity in distinct gut sections are still debated. In this study, we focused on the tropical millipede species Archispirostreptus gigas (Diplopoda, Spirostreptidae) and Epibolus pulchripes (Diplopoda, Pachybolidae), which release considerable amounts of methane. We aimed to characterize relationships between physicochemical parameters, methane production rates, and methanogen community structure in the two major gut sections, midgut and hindgut. Microsensor measurements revealed that both sections were strictly anoxic, with reducing conditions prevailing in both millipedes. Hydrogen concentration peaked in the anterior hindgut of E. pulchripes. In both species, the intestinal pH was significantly higher in the hindgut than in the midgut. An accumulation of acetate and formate in the gut indicated bacterial fermentation activities in the digestive tracts of both species. Phylogenetic analysis of 16S rRNA genes showed a prevalence of Methanobrevibacter spp. (Methanobacteriales), accompanied by a small fraction of so-far-unclassified “Methanomethylophilaceae” (Methanomassiliicoccales), in both species, which suggests that methanogenesis is mostly hydrogenotrophic. We conclude that anoxic conditions, negative redox potential, and bacterial production of hydrogen and formate promote gut colonization by methanogens. The higher activities of methanogens in the hindgut are explained by the higher pH of this compartment and their association with ciliates, which are restricted to this compartment and present an additional source of methanogenic substrates.
IMPORTANCE Methane (CH4) is the second most important atmospheric greenhouse gas after CO2 and is believed to account for 17% of global warming. Methanogens are a diverse group of archaea and can be found in various anoxic habitats, including digestive tracts of plant-feeding animals. Termites, cockroaches, the larvae of scarab beetles, and millipedes are the only arthropods known to host methanogens and emit large amounts of methane. Millipedes are ranked as the third most important detritivores after termites and earthworms, and they are considered keystone species in many terrestrial ecosystems. Both methane-producing and non-methane-emitting species of millipedes have been observed, but what limits their methanogenic potential is not known. In the present study, we show that physicochemical gut conditions and the distribution of symbiotic ciliates are important factors determining CH4 emission in millipedes. We also found close similarities to other methane-emitting arthropods, which might be associated with their similar plant-feeding habits.


In recent years, methane (CH4) has been at the center of discussion due to its profound contribution to global climate change (13). Most biogenic methane is produced by methanogens (for the exception, see reference 4), which are strictly anaerobic archaea and occur in almost all anoxic habitats on earth, including the digestive tracts of animals and humans (511). Among invertebrates, only termites, cockroaches, the larvae of scarab beetles, millipedes, one earthworm species, and marine copepods have been shown to host methanogenic microorganisms (1217). Termites have been estimated to contribute 1 to 3% of global methane production (18) and have been extensively studied with respect to methanogenesis (19).
Millipedes feed on decaying plant matter and are considered key species in various terrestrial ecosystems (20). As detritivores, millipedes depend on their intestinal microbiotas in the digestion of cellulose or recalcitrant humus (21). In general, methanogenesis may be driven by hydrogen, methylated C1 compounds, or acetate, and it represents the final step in the anaerobic degradation of lignocellulose (22, 23). Since an accumulation of hydrogen and acetate inhibits microbial fermentation processes, methanogenesis should exert positive feedback on symbiotic digestion in plant-feeding arthropods (24, 25).
Methane emission by millipedes was first observed in large tropical species (26). However, lower and more variable rates of methane production were also recorded in smaller European species (27). The factors limiting methanogenesis are not clear, but the activity of methanogens in millipede guts is correlated with a combination of factors such as host body size and geographical region (17, 27).
To date, little is known about the methanogen community in different sites of the millipede digestive tract. The digestive tract of millipedes is a straight tube consisting of a distinct foregut, midgut, pylorus, and hindgut (2831). In all arthropods studied, the hindgut represents the exclusive site of methane production (17, 3234). Methanogens can occur free-floating in the gut lumen, attached to the gut content or to the gut wall surface, or associated with anaerobic protists (26, 35, 36). Hackstein and Stumm (26) observed free-living as well as endosymbiotic methanogens in the hindguts of tropical millipedes, which were later identified as Methanobrevibacter-like methanogens (37). Paul et al. (38) enriched strain MpM2 from the hindgut of an Anadenobolus sp., which is a close relative of “Candidatus Methanoplasma termitum” (39) and represents a novel lineage of obligately methylotrophic methanogens (order Methanomassiliicoccales) that are widespread in the intestinal tracts of both invertebrate and vertebrate hosts. Šustr et al. (17) identified Methanosarcinales, Methanobacteriales, Methanomicrobiales, and other unclassified Archaea in the hindguts of various millipede species. While reducing conditions, a neutral pH, and the provision of H2 and other methanogenic substrates are considered prerequisites for colonization of the hindgut by methanogens (40, 41), the contributions of other factors remain to be explored.
In this study, we investigated the methane production and community structure of methanogens in the midgut and hindgut of the millipede species Archispirostreptus gigas and Epibolus pulchripes and determined physicochemical conditions limiting the methanogenic potential. A. gigas and E. pulchripes are large species from the orders Spirostreptida and Spirobolida, respectively, that are widely distributed in the tropics, hosting methanogens and releasing large amounts of methane (17).
We hypothesized that (i) in millipedes, the hindgut, rather than the midgut, would provide conditions favoring methanogens (anoxia, reducing conditions, high concentration of fermenting products, and presence of ciliates) and thus would represent a hot spot for methane production and (ii) the hindgut would possess a stable methanogenic community different from that in the midgut.


Methane emission rates.

Weight-specific methane emission rates were approximately two times higher in E. pulchripes (average, 17.4 nmol g of fresh wet weight−1 h−1) than in A. gigas (average, 6.5 nmol g of fresh wet weight−1 h−1) (Fig. 1A). Rates differed significantly between millipede species [t test: t(18) = −3.42; P = 0.003]. In both species, methane production in the hindgut was significantly higher than that in the midgut (Fig. 1B) [repeated-measures analysis of variance [ANOVA]: A. gigas, F(1,8) = 27.61, P = 0.001; E. pulchripes, F(1,8) = 30.55, P = 0.001].
FIG 1 (A) Methane emission rates of Archispirostreptus gigas (n = 10) and Epibolus pulchripes (n = 10) expressed in nanomoles per gram (fresh weight) of a millipede per hour. (B) Weight-specific methane emission rates of isolated midgut and hindgut sections of selected individuals (n = 5) measured at five time points (after 1, 2, 4, 6, and 24 h of anaerobic incubation). The methane production rates are expressed in nanomoles per gram (fresh weight) of a single gut section. All rates are reported as means ± SE. For original data on fresh weight of millipedes and gut sections, see Table S1.

Oxygen, redox, and pH conditions in the gut lumen.

Radial profiles of midguts and hindguts showed steep oxygen gradients toward the surface of freshly dissected guts of both species (Fig. 2). Oxygen concentration decreased to zero directly below the gut wall and remained below the detection limit along the entire radial profile inside both sections of the digestive tract in both A. gigas and E. pulchripes. The extent of oxygen penetration into the gut lumen was not resolved, but the microoxic periphery is restricted to a thin (<500-μm) layer below the gut wall. Axial profiles revealed that the gut lumen was largely anoxic in the whole volume of the midgut and hindgut (Fig. 2). The gut lumen of both species showed strongly reducing conditions (between −114 and −243 mV) along the entire intestinal tract. Redox potential was lowest in the posterior midgut (−242 and −243 mV for E. pulchripes and A. gigas, respectively) and increased steadily in the hindgut section (Table 1). The intestinal pH axial profiles showed similar courses in both species (Fig. 3A). Midgut pH was acidic and increased steadily from pH 4.4 to 6.1 in the posterior direction. Across the pyloric region (see Fig. S1 in the supplemental material), the pH increased to its maximum and remained slightly alkaline throughout the hindgut section (pH 7.3 to 7.9). The axial profile of H2 concentration in midgut and hindgut was measured in E. pulchripes only. The hydrogen level increased steadily in the posterior of the midgut, reached its maximum in the anterior part of the hindgut, and then decreased toward the posterior part (Fig. 3B).
FIG 2 Typical radial profiles of oxygen concentration in agarose-embedded midgut and hindgut sections of millipedes Archispirostreptus gigas and Epibolus pulchripes. “Depth” refers to the distance between the electrode tip and the surface of the agarose. To account for individual variations in the exact depths of embedding, the solid arrowheads indicate the points at which the tip reached the gut wall; the open arrowheads indicate the point of emergence on the opposite side. Readings were taken every 0.5 mm.
FIG 3 Axial profiles of pH and hydrogen concentration along the intestinal tracts of millipedes. (A) pH in A. gigas and E. pulchripes; (B) hydrogen concentration in E. pulchripes. Readings were taken at cardinal points of the respective gut sections.
TABLE 1 Redox potential of midgut and hindgut contents at the gut centers of Archispirostreptus gigas and Epibolus pulchripes
SpeciesRedox potential (mV) ina:
Archispirostreptus gigas−225 ± 51−242 ± 78−146 ± 68−114 ± 29
Epibolus pulchripes−155 ± 19−243 ± 33−212 ± 46−124 ± 45
All values are means ± SE for two individuals of A. gigas and five individuals of E. pulchripes.

Metabolic end products in the gut lumen and the hemolymph.

Acetate was the major fermentation product in the hindgut of both species. Propionate concentrations were noteworthy in the hindgut of A. gigas, whereas E. pulchripes accumulated only some formate (Fig. 4). E. pulchripes accumulated high concentrations of acetate and formate in the midgut, whereas short-chain fatty acid (SCFA) concentrations were quite low for A. gigas. The SCFA concentrations were generally low in the hemolymph, with detectable amounts of lactate, acetate, and succinate (Fig. 4).
FIG 4 Short-chain fatty acids in midgut, hindgut, and hemolymph of Archispirostreptus gigas (A) and Epibolus pulchripes (B). Concentrations are means ± SE for two and five individuals, respectively. Results are expressed in millimoles per liter of hemolymph or gut content volume.

Presence of ciliates and nematodes in hindgut.

The hindguts of both millipede species were densely colonized by large ciliates of the genus Nyctotherus and several species of nematodes. The average density of ciliates in the hindgut content was 1,041 ± 403 ml−1 (354 to 1,750 cells per individual) in A. gigas and 1,778 ± 424 ml−1 (933 to 2,267 cells per individual) in E. pulchripes. The average density of nematodes in the hindgut content was 386 ± 167 ml−1 in A. gigas and 711 ± 395 ml−1 in E. pulchripes. The nematodes were identified as Brumptaemilius sp., Abirovulva sp., and Rhigonema sp. by Vlastimil Baruš (personal communication). Neither protists nor nematodes were found in the midgut sections of both millipede species.

Methanogenic communities in the midgut, hindgut, and fecal pellets.

Clone libraries of the archaeal 16S rRNA genes from the different gut compartments and fecal pellets of the two millipedes yielded a total of 67 near-full-length sequences. Phylogenetic analysis revealed that both millipedes harbored members of the genus Methanobrevibacter (order Methanobacteriales) (Fig. 5) and so-far-unclassified members of the candidate family “Methanomethylophilaceae” (order Methanomassiliicoccales) (Fig. 6).
FIG 5 Phylogenetic tree (16S rRNA genes) of the genus Methanobrevibacter, illustrating the relationships of uncultured methanogenic archaea from the gut of Archispirostreptus gigas (green; 34 clones) and Epibolus pulchripes (red; 33 clones) to their closest relatives in public databases. Taxa that contain representatives from arthropod guts are in blue. The maximum-likelihood tree was reconstructed using IQ-TREE. Closed and open bullets indicate highly supported nodes with values of ≥95/99 and ≥75/90, respectively (SH-aLRT/ultrafast bootstrap analysis, 1,000 replicates each). Accession numbers are shown only for one representative per subgroup. For more details, including accession numbers of all sequences and outgroups, see Fig. S3.
FIG 6 Phylogenetic tree (16S rRNA genes) of the order Methanomassiliicoccales, showing the relationships of uncultured methanogenic archaea from the gut of Archispirostreptus gigas (green; 34 clones) and Epibolus pulchripes (red; 33 clones) to their closest relatives in public databases. Taxa that contain representatives from arthropod guts are in blue. The maximum-likelihood tree was reconstructed using IQ-TREE. Closed and open bullets indicate highly supported nodes with values of ≥95/99 and ≥75/90, respectively (SH-aLRT/ultrafast bootstrap analysis, 1,000 replicates each). Accession numbers are shown only for one representative per subgroup. For more details, including accession numbers of all sequences and outgroups, see Fig. S3.
Most Methanobrevibacter sequences fell into a monophyletic clade that included Methanobrevibacter arboriphilus. Except for a single clone from A. gigas, they represented distinct lineages (0.5 to 2% sequence dissimilarity [Table S2]) in the Mpn19 clade, which comprises the isolate Methanobrevibacter strain Mc30 (42) and several clones previously obtained from termites and scarab beetle larvae. In addition, each millipede harbored a novel lineage that does not group with any isolates and is sufficiently distinct (3 to 5% sequence dissimilarity [Table S2]) from the clones in the M. arboriphilus group to qualify as a separate species. In the case of E. pulchripes, the clones clustered with uncultured phylotypes from termite guts (P4b-Ar-15 group); in the case of A. gigas, they formed a deep-branching clade (Ag101-11 group) that is well separated from all other species groups (Methanobrevibacter smithii and Methanobrevibacter ruminantium groups) (Fig. 5).
The sequences that grouped with the “Methanomethylophilaceae” were recovered only from a few of the libraries and always in small numbers (Table S2). The phylotypes were quite similar to each other (<1% sequence dissimilarity) (Table S2) and represented three distinct host-specific lineages that clustered with uncultured methanogens from the hindgut of a scarab beetle larva (PeHAr25 group) (Fig. 6).



Methane emission by the midguts of A. gigas and E. pulchripes was only marginal, confirming that methanogenesis in arthropods is generally restricted to the hindgut (12). The weight-specific methane emission rates of millipedes (10 to 100 nmol g−1 h−1) are in the same range as those of cockroaches and scarab beetle larvae (26, 27, 43) but 1 order of magnitude lower than those of termites (4446). The proximity of H2-reducing and H2-consuming gut sections in the abdomen of termites and cockroaches is considered an important prerequisite for methanogenesis (34, 47, 48). However, the intestinal tract of millipedes is a straight tube without any direct contact between midgut and hindgut sections (Fig. S1), which precludes any transepithelial H2 exchange. It remains possible that less volatile substrates (i.e., formate and methanol) are transported from midgut to hindgut with the digesta or via the hemolymph (12).

Physicochemical gut conditions.

As in all other arthropods studied to date (for example, see references 41, 48, 49, and 50), the lumen of the midgut and hindgut compartments of A. gigas and E. pulchripes is anoxic. Anoxia of the gut contents of invertebrates is attributed to the oxygen consumption of strictly and facultatively aerobic bacteria located in the peripheral region of the gut lumen (51, 52). Also, millipede guts are densely colonized by microorganisms (21, 53), which might serve as an oxygen sink.
Methanogens require not only anoxia but also a negative redox potential (<−100 mV) (54, 55). This agrees with the reports of reducing conditions in the hindguts of all methane-emitting arthropods (40, 41, 50, 5658) and their absence from the hindguts of species that do not emit methane (57, 59, 60). The gut environment of A. gigas and E. pulchripes provides strongly reducing conditions (from −114 to –243 mV) not only in the hindgut but also in the midgut compartment, which emits only traces of methane. This resembles the situation in cockroaches, where the redox potential in the midgut is often more negative than in the hindgut, while methanogenesis is restricted to the hindgut compartment (40, 50). In termites and methane-emitting coleopteran larvae, the redox potential in the midgut is typically higher (less negative or positive) than in the hindgut (41, 5658) and therefore unfavorable for methanogens.
Another environmental factor that may influence methanogenesis in the gut of millipedes is the intestinal pH. All hindgut compartments of methane-emitting insects studied to date are either neutral or slightly alkaline (40, 61, 62), and in the rumen, an acidic pH inhibits methanogenesis (63). Thus, the slightly alkaline pH in the hindgut of millipedes may be more favorable for methanogens than the acidic conditions in the midgut, explaining the low rates of methanogenesis observed in this compartment. However, the slightly to highly alkaline pH in the midgut of scarab beetle larvae indicates that there must be other parameters that exclude methanogens from the midgut compartment (41, 58).

Fermentation products in gut sections and methanogenesis.

The presence of SCFAs in insect guts indicates bacterial fermentations (for example, see references 41, 56, and 64). As in other invertebrates (for example, see references 24, 65, and 66), acetate is the dominant SCFA in both gut sections of A. gigas and E. pulchripes. The acetate concentration was lower than in wood-feeding termites (66) or cockroaches (64) but similar to that in soil-feeding termites (67). Propionate, another typical fermentation product in the hindguts of insects (41, 50, 66, 68), was the second most abundant SCFA in the hindgut of A. gigas. Lactate, which is typically present in insect hemolymph (41, 64, 66), was detected in low concentrations in both millipedes.
The guts of both A. gigas and E. pulchripes were densely colonized by parasitic nematodes, which are commonly encountered in the intestinal tracts of millipedes (for example, see references 69 and 70). Many nematodes from anoxic gut ecosystems are capable of fumarate respiration and form large amounts of succinate, which is metabolized to propionate by gut bacteria (71, 72). This would explain why succinate concentrations were rather low in both millipedes but propionate accumulated in the hindgut compartments of A. gigas, the species with the higher density of nematodes.
Methanogenesis in arthropod guts is generally hydrogenotrophic or methylotrophic; as in other intestinal systems, there is no evidence for acetoclastic methanogenesis (12, 19). Hydrogen is by far the most prominent substrate of methanogenesis in the hindguts of insects (for example, see references 34 and 73). In cockroaches and lower termites, most of the H2 is produced by the fermentative activities of anaerobic protists (32, 7476), whereas in higher termites, H2 is formed exclusively by the bacterial gut microbiota (48, 77, 78).
Like other millipede species (26), both A. gigas and E. pulchripes host large numbers of Nyctotherus-like ciliates in their hindguts (Fig. S1) (40). These protists possess hydrogenosomes and are usually associated with endosymbiotic methanogens (79). Nevertheless, the accumulation of hydrogen in the hindgut of E. pulchripes indicates that its production by ciliates and/or fermenting bacteria and methanogenesis might not be tightly coupled.
Formate was detected in both gut compartments of A. gigas and E. pulchripes. It is an important substrate for hydrogenotrophic methanogenesis in soil-feeding termites (48) and stimulates methanogenesis in isolated hindguts of cockroaches (32) and scarab beetle larvae (41). Although methanol was not analyzed in this study, it may be derived from microbial degradation of plant material and utilized by methylotrophic methanogens, as demonstrated for cockroaches (8082).

Methanogen diversity and community composition.

All methanogenic archaea identified in the guts of A. gigas and E. pulchripes are most closely related to representatives from the guts of other arthropods. With the exclusive presence of Methanobacteriales and only a few Methanomassiliicoccales, the archaeal community resembles that of scarab beetle larvae (58, 83) and cockroaches (84).
The Methanobacteriales are represented exclusively by members of the genus Methanobrevibacter, the most common genus of methanogens found also in the gut of insects (for a review, see reference 12). Each millipede harbors several host-specific lineages that fall into the Methanobrevibacter arboriphilus group. Although the clade MPn19 is distinct from the lineage comprising the type strain originally isolated from diseased trees (85) and consists exclusively of representatives associated with the intestinal tracts of arthropods, the high level of sequence similarity indicates that they may represent the same species (Fig. 5; Table S2). In contrast, the host-specific lineages in the P4b-Ar-15 group (E. pulchripes) and the Ag101-11 group (A. gigas) are well separated from all described Methanobrevibacter species and undoubtedly represent novel species-level lineages of so-far-uncultured methanogens from arthropod guts.
Members of the genus Methanobrevibacter are hydrogenotrophic, and the isolates from termite guts produce methane from H2/CO2 and sometimes from formate (73, 86). Many of the endosymbiotic methanogens found to be associated with hydrogen-producing Nyctotherus ciliates (37) grouped with the M. arboriphilus group, particularly the MPn19 clade (details not shown). Interestingly, the Mpn19 clade also comprises the reportedly formicotrophic Methanobrevibacter strain Mc30 (42), which suggests that the formate accumulating in the midgut and hindgut compartments contributes to methanogenesis in both millipedes. In contrast, all members of the “Methanomethylophilaceae” (order Methanomassiliicoccales), which are typically found in the intestinal tracts of both invertebrate and vertebrate hosts (38, 87), possess an obligately hydrogen-dependent methylotrophic metabolism (39, 88). It is therefore likely that methanogens of the PeHAr25 group, which are present in both millipedes, albeit in smaller numbers, utilize methanol derived from polysaccharide fermentation.


Methanogenesis, as the final step of polysaccharide degradation in the hindgut of millipedes, is driven by a complex interplay of abiotic and biotic factors. As in other arthropods, it is favored by anoxia, reducing conditions, and the neutral pH of the gut environment. Methanogenesis in millipedes is catalyzed by hydrogenotrophic Methanobrevibacter spp. and to some extend also methylotrophic members of Methanomassiliicoccales. The association of hydrogenotrophic methanogens with the abundant population of anaerobic ciliates should contribute significantly to methane production. The composition of the methanogenic community in millipedes is similar to that of scarab beetle larvae and comprises several species-level lineages of so-far-uncultured methanogens that are exclusively associated with the intestinal tracts of arthropods.



Archispirostreptus gigas and Epibolus pulchripes were reared at the Institute of Soil Biology (Biology Centre CAS, Czech Republic) for several years (for example, see reference 17). Animals were kept in plastic boxes (60 cm by 30 cm by 20 cm [height]) filled with a mixture of horticultural substrate and hardwood leaf litter. The substrate was moistened regularly with tap water. Millipedes were provided with their preferred food: E. pulchripes was fed with decomposed litter of maple and oak leaves, and A. gigas was offered fresh vegetables and fruits besides the leaf litter. Cuttlefish bone powder was added to the substrate as a source of calcium. Animals were kept at a constant temperature of 25°C. Only adult individuals were used for the experiments.

CH4 production rates.

Methane production was measured by placing individual millipedes into 100-ml glass bottles. Each millipede was weighed before methane measurements (n = 10 per species) (Table S1). Filter paper soaked with distilled water was placed in each bottle to maintain the air humidity inside the bottle, and bottles were sealed with rubber stoppers. The bottles were incubated for 4 h at 20°C in the dark. Samples (0.5 ml) of the internal atmosphere were collected at the start and at the end of the incubation period and immediately injected into a gas chromatograph inlet system for analysis (HP 5890 series II; Hewlett Packard, Palo Alto, CA, USA). The small changes of methane concentration inside control bottles, subjected to the same handling but without animals, were subtracted. Immediately after the measurements, five randomly selected individuals of each species were dissected, and the midgut and hindgut sections were separated (the midgut-hindgut interface is clearly visible [Fig. S1]), weighed (Table S1), and each placed into a 100-ml glass bottle containing 2 ml of Ringer’s solution (NaCl, 7 g liter−1; KCl, 0.25 g liter−1; CaCl2, 3 g liter−1). The headspace of the bottles was exchanged against an argon atmosphere. Samples of 0.5 ml of headspace gas were taken and analyzed for methane production after 1, 2, 4, 6, and 24 h of incubation, as described for the intact animals.

Microsensor measurements.

Oxygen concentration, redox potential, pH, and hydrogen concentration in the millipede gut were determined with microsensors (as described in references 47, 49, and 59). Oxygen concentration in gut content was determined with a Unisense microsensor monometer using a miniaturized Clark-type oxygen sensor with a guard cathode (OX-25; tip diameter, 20 to 30 μm; Unisense A/S, Denmark). The microsensor was calibrated by the two-point method. The air-saturated water was used for the atmospheric reading. Oxygen concentration in the saturated water was calculated using the Ostwald solubility coefficient (89) and adjusted to the temperature and atmospheric pressure of the experiment. A 20 g liter−1 solution of sodium ascorbate in 0.1 M NaOH was used for the zero point.
Redox potential (Eh) was measured with a Unisense pH/mV meter with a combination of the redox microelectrode RD-25 and the reference electrode REF-25 (tip diameters of 20 to 30 μm). For calibration, we used freshly prepared solutions of 10 g liter−1 quinhydrone in Britton-Robinson buffers of pH 4 and pH 7 as recommended by the microsensor manufacturer. Two components of the Britton-Robinson buffer A and buffer B were mixed in a volumetric ratio 4:1 or 1.75:1 for pH 4 and 7, respectively.
After dissection, the intact guts (Fig. S1) were immediately placed in a small chamber (200 mm by 23 mm by 35 mm high) on the surface of a 7- to 8-mm-thick layer of 7% agar made up with insect Ringer’s solution. Guts were quickly covered with a layer of 3% agar (temperature of 35°C), which cooled and solidified immediately. Microsensors were positioned vertically above the chamber with manual micromanipulators. For axial profiles of oxygen concentration and redox potential, measurements were routinely made in the anterior and posterior midgut and hindgut. Radial profiles of oxygen concentration were measured at the same positions with 0.5-mm increments. Each set of measurements was conducted independently for five individuals of E. pulchripes and two individuals of A. gigas.
Glass pH microelectrodes with a tip diameter of 20 to 30 μm (90), in combination with the reference electrode REF-25, were used to measure axial profiles of intestinal pH. They were calibrated with commercial pH standard solutions (pH 5, 6, 7, and 10). Measurements of axial profile of intestinal hydrogen concentration were made only in E. pulchripes, using H2 microsensors with a tip diameter of 20 to 30 μm. They were calibrated using two-point calibration (0 and 5% [by volume] H2).
After calibration, millipedes were dissected, and the intact gut was immediately placed in the measuring chamber (Fig. S1, same as above). The pH was measured in the center of the gut lumen in points proportionally distributed along the gut longitudinal axis of E. pulchripes and A. gigas. The measurements were conducted in three individuals of E. pulchripes and two individuals of A. gigas. All physicochemical parameters were measured at room temperature.

Metabolites in the gut fluid.

Concentrations of short-chain fatty acids (SCFA) were determined for midgut and hindgut sections and for the hemolymph of both species. Before dissection, the animals were immobilized by cooling, and 20 to 30 μl hemolymph was collected by intersegmental puncture at the dorsal part of the body using a Hamilton syringe. The intestinal contents (100 mg of fresh weight) and the collected hemolymph were homogenized and centrifuged at 15,000 × g for 10 min at 5°C. The supernatants were immediately used for high-performance liquid chromatography (HPLC) measurements. Low-molecular-weight organic acids were determined by analytical ion chromatography (Integrion, Thermo Fisher, USA) equipped with IonPac AG11-HC 4-μm and IonPac AS11-HC 4-μm columns (Thermo Fisher, USA) with conductivity detection. For elution, high-pressure analytical eluent generator KOH cartridges with multistep nonlinear gradient at a concentration from 1 mM to 85 mM were used. Analytical flow was 0.38 ml min−1. Samples were injected with a cooled (5°C) Dionex AS-AP autosampler (Thermo Fisher, USA) using partial-loop injection mode. The volume of injection for standards (Sigma-Aldrich) and samples was 15 μl, with a very low draw speed of the syringe. Measurements were replicated in five individuals of E. pulchripes and two individuals of A. gigas.

Ciliate protists and nematodes in the gut content.

The presence of large ciliates of the genus Nyctotherus (Fig. S2) and nematodes (Movie S1) in the midgut and the hindgut of both millipede species was verified directly with a binocular magnifier. Only living ciliates and nematodes were enumerated using a droplet method. Ciliates of Nyctotherus spp., which are commonly associated with endosymbiotic methanogens in tropical millipedes (26), can be easily distinguished by their distinct morphology and movement. After dissection (as described above), an aliquot of the midgut and hindgut content was diluted using tap water (1:4 [vol/vol]), and a drop (10 μl) was spread on the microscopic slide. All living nematodes and large Nyctotherus ciliates were counted in each 12 replicates. Hindgut contents of three individuals of A. gigas and three individuals of E. pulchripes were examined.

DNA extraction and purification.

Three individuals of A. gigas and four individuals of E. pulchripes species were transferred to sterile petri dishes, and fecal pellets were continuously collected in Eppendorf tubes for several hours. Millipedes were anaesthetized by cold (−20°C) and killed by decapitation. Individuals were dissected under sterile conditions, and guts were separated into midgut and hindgut sections. Gut sections and fecal pellets were frozen immediately. Total genomic DNA was extracted using a NucleoSpin Tissue XS kit (Macherey-Nagel, Germany). Tissue samples (10 mg) were digested with proteinase K overnight at room temperature. Extractions were performed according to the manufacturer's instructions. The quality of extracted DNA was verified by standard gel electrophoresis, and DNA was quantified with a NanoDrop spectrophotometer (Thermo Scientific, Wilmington, DE, USA).

16S rRNA gene libraries and phylogenetic analysis.

16S rRNA gene libraries of Archaea were constructed with primers Ar109f and 1492 (91, 92). Each PCR mixture (50 μl) contained reaction buffer, 2.5 mM MgCl2, 1 U Taq DNA polymerase (all from Invitrogen, Carlsbad, CA, USA), a mixture of deoxynucleoside triphosphates (dNTPs) at 50 μM, a 0.3 μM concentration of each primer, 0.8 mg ml−1 bovine serum albumin (BSA; Thermo Scientific, Waltham, MA, USA), and 1 μl template DNA. The PCR program consisted of an initial denaturation step (94°C for 3 min), followed by 32 cycles of denaturation (94°C for 20 s), annealing (52°C for 20 s), and extension (72°C for 50 s), and a final extension step (72°C for 7 min). PCR products were purified using the MinElute PCR purification kit (Qiagen, Hilden, Germany). The amplified products of archaeal 16S rRNA genes were cloned into pGEM-T Easy vector (Promega Corporation, Madison, WI, USA). Selected clones (blue/white selection) were screened for the correct insert size by gel electrophoresis, and inserts were amplified using primer M13 (Biotech, Germany) and commercially sequenced.
For phylogenetic analysis, the sequences were imported into the reference alignment of the Silva SSURef database (version 132; (93) using the ARB software package (version 6.0.6) (94), including additional sequences retrieved from GenBank ( The alignment was manually refined with the ARB editor, using the secondary structure of the rRNA to identify homologous base positions. For each library, potentially chimeric sequences were identified by fractional treeing and by inspection of the signature sequences of the respective clades. Chimeric sequences were excluded from the analysis (Table S2). After removing sites with >50% gaps, the alignment consisted of 1,325 sites with unambiguously aligned base positions, of which 288 were invariant and 863 were parsimony-informative sites. Phylogenetic trees were reconstructed by maximum-likelihood analysis with IQ-TREE (95) using the best-fit evolutionary model (SYM+I+G4) suggested by ModelFinder (96). Node support was assessed with the Shimodaira-Hasegawa approximate likelihood ratio test (SH-aLRT) (97) and by ultrafast bootstrap analysis (UFBoot [98]). Trees were visualized in ARB and polished in Inkscape (

Statistical analyses.

Unless mentioned otherwise, data are presented as means and standard errors (SE) of the results obtained with at least four different animals. Differences in weight-specific methane production rates between species were evaluated by t test. Differences in methane production rates between midgut and hindgut sections were analyzed separately for each species using repeated-measures analysis of variance (ANOVA). Statistical analyses were performed with STATISTICA version 6.0 (Statsoft Inc., USA).

Data availability.

Sequencing reads were submitted to GenBank under accession numbers MN722465 to MN722531.


This work was supported by the Czech Science Foundation (project no. 17-22572S), the Grant Agency of the University of South Bohemia (project no. 04-145/2010/P), the Ministry of Education, Youth and Sports of the Czech Republic (project no. LM2015075 and EF16_013/0001782), Charles University Research Centre program no. 204069, the Collaborative Research Center SFB 987 of the German Research Foundation (DFG), and the Max Planck Society (MPG). T.H. was supported by a PPPLZ grant from the Czech Academy of Sciences (L200961851).
We thank Linda Jíšová and Lucie Faktorová for technical assistance, Marie Stehlíková for help with laboratory analyses, Vlastimil Baruš for the identification of nematodes, and Aleksandra Walczyńska for commenting on the manuscript.

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Published In

cover image Applied and Environmental Microbiology
Applied and Environmental Microbiology
Volume 87Number 1513 July 2021
eLocator: e00614-21
Editor: Harold L. Drake, University of Bayreuth
PubMed: 34020937


Received: 31 March 2021
Accepted: 12 May 2021
Accepted manuscript posted online: 21 May 2021
Published online: 13 July 2021


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  1. methane
  2. methanogenic community
  3. Methanobrevibacter
  4. physicochemical parameters
  5. digestive tract
  6. Methanomassiliicoccales
  7. methanogenesis
  8. tropical millipedes



Institute of Soil Biology, Biology Centre CAS, České Budějovice, Czech Republic
Present address: Terézia Horváthová, EAWAG, Swiss Federal Institute of Aquatic Science and Technology, Dübendorf, Switzerland.
Vladimír Šustr
Institute of Soil Biology, Biology Centre CAS, České Budějovice, Czech Republic
SoWa Research Infrastructure, Biology Centre CAS, České Budějovice, Czech Republic
Alica Chroňáková
Institute of Soil Biology, Biology Centre CAS, České Budějovice, Czech Republic
SoWa Research Infrastructure, Biology Centre CAS, České Budějovice, Czech Republic
Stanislava Semanová
Institute of Soil Biology, Biology Centre CAS, České Budějovice, Czech Republic
Faculty of Science, University of South Bohemia, České Budějovice, Czech Republic
Kristina Lang
RG Insect Gut Microbiology and Symbiosis, Max Planck Institute for Terrestrial Microbiology, Marburg, Germany
Carsten Dietrich
RG Insect Gut Microbiology and Symbiosis, Max Planck Institute for Terrestrial Microbiology, Marburg, Germany
Tomáš Hubáček
SoWa Research Infrastructure, Biology Centre CAS, České Budějovice, Czech Republic
Masoud M. Ardestani
SoWa Research Infrastructure, Biology Centre CAS, České Budějovice, Czech Republic
Institute for Environmental Studies, Charles University in Prague, Prague, Czech Republic
Ana C. Lara
Institute of Soil Biology, Biology Centre CAS, České Budějovice, Czech Republic
Andreas Brune
RG Insect Gut Microbiology and Symbiosis, Max Planck Institute for Terrestrial Microbiology, Marburg, Germany
Miloslav Šimek
Institute of Soil Biology, Biology Centre CAS, České Budějovice, Czech Republic
Faculty of Science, University of South Bohemia, České Budějovice, Czech Republic


Harold L. Drake
University of Bayreuth

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