Bacteria and archaea involved in anaerobic digestion ... - Springer Link

Appl Microbiol Biotechnol (2011) 89:2039–2052 DOI 10.1007/s00253-010-2981-9

ENVIRONMENTAL BIOTECHNOLOGY

Bacteria and archaea involved in anaerobic digestion of distillers grains with solubles Ayrat M. Ziganshin & Thomas Schmidt & Frank Scholwin & Olga N. Il’inskaya & Hauke Harms & Sabine Kleinsteuber

Received: 27 September 2010 / Revised: 20 October 2010 / Accepted: 20 October 2010 / Published online: 9 November 2010 # Springer-Verlag 2010

Abstract Cereal distillers grains, a by-product from bioethanol industry, proved to be a suitable feedstock for biogas production in laboratory scale anaerobic digesters. Five continuously stirred tank reactors were run under constant conditions and monitored for biogas production and composition along with other process parameters. Iron additives for sulfide precipitation significantly improved the process stability and efficiency, whereas aerobic pretreatment of the grains had no effect. The microbial communities in the reactors were investigated for their phylogenetic composition by terminal restriction fragment length polymorphism analysis and sequencing of 16S rRNA genes. The bacterial subcommunities were highly diverse, and their composition did not show any correlation with reactor performance. The dominant phylotypes were affiliated to the Bacteroidetes. The archaeal subcommunities were less diverse and correlated Electronic supplementary material The online version of this article (doi:10.1007/s00253-010-2981-9) contains supplementary material, which is available to authorized users. A. M. Ziganshin : O. N. Il’inskaya Department of Microbiology, Kazan (Volga Region) Federal University, ul. Kremlevskaya 18, 420008 Kazan, The Republic of Tatarstan, Russian Federation T. Schmidt : F. Scholwin Department of Biochemical Conversion, German Biomass Research Centre, Torgauer Str. 116, 04347 Leipzig, Germany H. Harms : S. Kleinsteuber (*) Department of Environmental Microbiology, UFZ—Helmholtz Centre for Environmental Research, Permoserstr. 15, 04318 Leipzig, Germany e-mail: [email protected]

with the reactor performance. The well-performing reactors operated at lower organic loading rates and amended with iron chloride were dominated by aceticlastic methanogens of the genus Methanosaeta. The well-performing reactor operated at a high organic loading rate and supplemented with iron hydroxide was dominated by Methanosarcina ssp. The reactor without iron additives was characterized by propionate and acetate accumulation and high hydrogen sulfide content and was dominated by hydrogenotrophic methanogens of the genus Methanoculleus. Keywords Biogas . DDGS . T-RFLP . Methanogenic archaea . Porphyromonadaceae . Actinomycetales

Introduction Biomass is a readily available renewable energy source that has received increasing attention due to rising prices of fossil fuels and the urgent need to mitigate anthropogenic global warming. The conversion of biomass into gaseous and liquid biofuels by microorganisms can be considered as a way to gain safe and sustainable energy that does not contribute to a further buildup of carbon dioxide in the atmosphere (McKendry 2002). While the use of energy crops for bioenergy production leads to increase in food costs, the utilization of agricultural by-products and residues for the production of biofuels, especially biogas, can increase the economic and ecological benefit of biomass use and concomitantly prevent the accumulation of harmful biowaste in the environment (Ahn et al. 2009). Generation of biogas from biomass is driven by microbes, therefore, the microbial communities involved in this process and the impact of operational conditions on their activities received increasing attention during the last

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years (Antoni et al. 2007; Narihiro and Sekiguchi 2007; Chen et al. 2008; Demirel and Scherer 2008; Weiland 2010). Whereas the microbiology of anaerobic digesters for wastewater treatment is well investigated (Narihiro and Sekiguchi 2007; Chen et al. 2008), knowledge on the biological catalysts in pure plant biomass fermenters is still scarce (Antoni et al. 2007). The studies published so far focused mostly on the final stages of anaerobic digestion— the aceticlastic and hydrogenotrophic methanogenesis—and thus addressed mesophilic and thermophilic archaea in lab- or plant-scale bioreactors containing various agricultural and municipal wastes (Klocke et al. 2007; Demirel and Scherer 2008; Klocke et al. 2008; Nettmann et al. 2008; Weiss et al. 2009; Goberna et al. 2009, 2010; Lee et al. 2009; Krakat et al. 2010). Methanomicrobiales, Methanobacteriales, Methanococcales, and Methanosarcinales are the four phylogenetic orders of methanogens that are currently known to be present in mesophilic anaerobic digesters (Demirel and Scherer 2008). Aceticlastic methanogenesis is exclusively performed by members of the genera Methanosaeta and Methanosarcina, whereas other species of the Methanosarcinales are methylotrophs and some can also use hydrogen and carbon dioxide (Kendall and Boone 2006). Members of the other three orders use hydrogen and carbon dioxide or formate for growth and produce methane as gaseous waste. Functional groups of methanogens can be distinguished based on their phylogenetic affiliation, thus rRNA-targeting probes for FISH (Karakashev et al. 2005; Krakat et al. 2010) or microarrays (Franke-Whittle et al. 2009; Goberna et al. 2010) are applicable to monitor the methanogenic pathways in anaerobic digesters. In contrast to the studies on methanogenic archaea, the body of literature describing bacterial diversity in mesophilic agricultural biogas plants is small (O’Sullivan et al. 2005; Cirne et al. 2007; Klocke et al. 2007; Krause et al. 2008; Kröber et al. 2009; Liu et al. 2009). The initial stages of anaerobic digestion, in particular hydrolysis and acidogenesis, are brought about by complex bacterial communities (primary fermenters) that vary depending on substrate, fermenter type (e.g., single-stage vs. two-stage), and process conditions (e.g., mesophilic vs. thermophilic; Antoni et al. 2007). Depending on feedstock composition and process conditions, the hydrolysis of complex organic material by bacterial exoenzymes might be the rate-limiting step in anaerobic digestion of plant biomass (Lübken et al. 2007). Thus, to increase the efficiency of methane formation from plant biomass, the hydrolysis step needs to be enhanced. One way to improve the hydrolysis efficiency is inoculation of bacteria with high hydrolytic activities into bioreactors (Bagi et al. 2007; Yang et al. 2009). For some substrates, it is essential to apply appropriate inocula for effective anaerobic degradation (Angelidaki et al. 2002). Further measures to enhance

Appl Microbiol Biotechnol (2011) 89:2039–2052

hydrolysis efficiency are the separation of the hydrolysis/ acidogenesis stage from the acetogenesis/methanogenesis stage or the application of thermophilic processes (Antoni et al. 2007). However, the more diverse and recalcitrant feedstock for biogas production becomes, the more important is the exploitation of novel biocatalysts with higher hydrolytic activities (Alper and Stephanopoulos 2009). Thus, a more detailed understanding of the functional diversity of primary fermenters in biogas reactors is a prerequisite for knowledge-based process optimization. Some of the fermentation products of acidogenesis such as hydrogen, carbon dioxide, acetate, and formate—can be directly used by aceticlastic and hydrogenotrophic methanogens. Other fermentation products—mainly volatile fatty acids (VFA) and alcohols—are further converted to acetic acid, carbon dioxide, and hydrogen by acetogenic bacteria. The acetogenesis involves syntrophic secondary fermenters as well as homoacetogens (Schink 1997) and has also been regarded as the rate-limiting step of biogas formation (Antoni et al. 2007). Due to their dependency on interspecies hydrogen transfer, syntrophic secondary fermenters are most severely affected by a failure in methanogenic hydrogen and formate removal, e.g., due to a drop in pH or the presence of toxic compounds (Schink 1997). Sulfate-reducing bacteria (SRB) are also present in anaerobic digesters in relative abundances depending on the sulfate load. They compete with methanogens, acetogens, and fermenting bacteria for hydrogen, acetate, and other organic substrates. Moreover, H2S produced by SRB inhibits various bacterial groups as well as methanogens (Chen et al. 2008). Detailed analysis of community composition and dynamics is needed to unravel the complex antagonistic and synergistic effects in anaerobic digester communities and eventually improve process stability and efficiency of biogas formation. Molecular techniques targeting phylogenetic or functional marker genes have unveiled the complexity of microbial communities in anaerobic digesters and revealed the presence of not yet cultivable microorganisms (Narihiro and Sekiguchi 2007; Talbot et al. 2008). In the present study, molecular methods targeting bacterial and archaeal 16S rRNA genes as phylogenetic markers were used to analyze bacteria and archaea in mesophilic lab-scale digesters with distillers dried grains with solubles (DDGS) as a substrate. DDGS tested in our research is a dried by-product of ethanol distilling gained from grain’s mash. Usually, DDGS and wet distillers grains with solubles (WDGS) are used for animal nutrition. To increase shelf life and reduce transportation costs, wet distillers grains have to be dried, which is energy-intensive and consumes about one third of the energy demand of an entire dry grind plant (Bothast and Schlicher 2005). Moreover, the use of WDGS or DDGS as cattle feed


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requires high quality standards. Therefore, anaerobic digestion of WDGS to biogas could be an alternative source of revenue for dry grind ethanol production plants. In addition, microorganisms involved in distillers grains anaerobic utilization coupled with biogas production are not yet described in the literature. Here, we tested cereal distillers grains with solubles as a possible substrate for an anaerobic digestion process. Furthermore, the effects of H2S removal and substrate pretreatment on the microbial community structure and performance of the biogas reactors were investigated.

Materials and methods Digester configurations and running conditions Five continuously 7-L stirred tank reactors of similar design with 5 L working volume were fed with distillers grains (DDGS; CropEnergies AG, Zeitz, Germany) and operated at 38°C (Table 1). During set up, all digesters were inoculated with 20 mL outflow from another fermenter also fed with DDGS. The digester no. 6.2 was then fed only with DDGS and water. The feedstock for the digester no. 6.3 was composed of DDGS, water, and FerroSorp® DG, a powdery desulfurization substance containing ferric hydroxide (HeGo, Biotec, Germany). Before its addition to the digester, a 250-mL Erlenmeyer flask containing 30.6 g DDGS, 118.18 mL water, and 20 g of the outflow from the same fermenter was aerobically incubated on a rotary shaker at 100 rpm and room temperature for 2 days. Pretreatment of DDGS with the fermenter effluent was assayed to see whether the fermenter microflora would increase the rate of substrate hydrolysis under aerobic conditions. The pretreated substrate, additional 50 mL of water, and 1.22 g of FerroSorp® DG were transferred to the bioreactor no. 6.3. FerroSorp® DG was used as an agent to remove H2S from biogas.

The feedstock for the digester no. 6.4 was prepared as follows: 200 g of DDGS was added to 1.0 L of water, the mixture was then mixed well and centrifuged at 4,500×g for 3 min, and 102.4 g of the liquid phase, containing also large floating particles, was used as the substrate. Pretreatment of the substrate with the fermenter outflow (20 g) for 2 days was carried out as well. The pretreated substrate was transferred to the fermenter no. 6.4, accompanied by the addition of 46.21 mL of water and 1.39 g of 40% FeCl3. FeCl3 was used as an additive to precipitate H2S in the form of iron sulfide from biogas in the digesters with lower organic loading rates (OLR). The feedstock for the digester no. 6.5 was composed of 12.24 g DDGS, 136.37 mL water, 20 g of the fermenter outflow, and 1.39 g of 40% FeCl3. Pretreatment of the DDGS with the fermenter outflow in 86.37 mL water was done accordingly, and 50 mL additional water and FeCl3 were added to the fermenter jointly with the treated DDGS. The feedstock for the digester no. 6.6 was the same as for the digester no. 6.5 with the exception that the feedstock was transferred to the digester without pretreatment. The amounts of DDGS and other components that were added to the bioreactors are summarized in Table 1. All bioreactors were fed every day, and digested DDGS was also withdrawn daily. Volume and composition of the generated biogas as well as pH of the digestion mixtures were analyzed every day, while acid capacity and concentrations of ammonium and VFA were measured twice a week. Analytical methods Biogas volume was monitored using a milligascounter Ritter MGC-1 (Bochum, Germany). Biogas composition was measured by an infrared landfill gas analyzer GA 94 (Ansyco, Karlsruhe, Germany). Ammonium levels were analyzed in the liquid phase of the samples after removal of the solid phase by centrifugation at 20,000×g for 20 min. The supernatant was colored with Nessler’s reagent (Merck,

Table 1 Digesters’ configurations and running conditions TS oTS Inlet (gday−1) Digester Volume of Operating Organic no. digesting temperature loading rate (%) (%) mixture (L) (°C) (goTSday−1) DDGS H2O

Hydraulic retention Inoculum FerroSorp® DG FeCl3 (40%) Sum time (day)

6.2 6.3 6.4

5.0 5.0 5.0

38 38 38

10 25 10

88 88 11

81.8 12.24 137.76 – 81.8 30.6 168.18 20 9.8 102.4a 46.21 20

– 1.22 –

– – 1.39

150 220 170

33.3 22.7 29.4

6.5 6.6

5.0 5.0

38 38

10 10

88 88

81.8 12.24 81.8 12.24

– –

1.39 1.39

170 150

29.4 33.3

136.37 20 136.37 –

oTS organic total solids, TS (%) total solids (%) in DDGS, oTS (%) organic total solids (%) in DDGS a

The substrate for the digester no. 6.4 was prepared as follows: 200 g DDGS were added to 1.0 L water, the mixture was then mixed well and centrifuged at 4,500×g for 3 min, and 102.4 g of the liquid phase, containing not sedimented particles as well, were used as the substrate

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Darmstadt, Germany) and measured at 425 nm using a DR/ 2000 spectrophotometer (Hach Company, Loveland, USA). Acid capacity of the supernatants was determined by titration with 0.025–0.1 M H2SO4 in a pH range from 4.5 to 3.5 (Sawyer et al. 1994) using a Titration Excellence T90 titrator (Mettler-Toledo, Switzerland). Concentrations of VFA were determined by using a 5890 series II GC (Hewlett Packard, USA) equipped with an HS40 automatic headspace sampler (Perkin Elmer, USA) and an Agilent HP-FFAP column (30 m×0.32 mm×0.25 μm). The carrier gas was N2 (17.0 mLmin−1), and the oven temperature was raised firstly from 60°C to 100°C at a rate of 20°Cmin−1, then to 140°C at 5°Cmin−1, and finally to 200°C at 40°Cmin−1. The temperatures of injector and detector were 220°C and 250°C, respectively. Samples were analyzed after transfer of 0.5 mL of 85% H3PO4 into 10-mL vials containing 3.0 mL of the supernatants. The vials were then closed with caps, and the gaseous phase was injected into the GC. Acetate, propionate, isobutyrate, and other organic acids (purchased from SigmaAldrich, Germany) were used as standards. DNA extraction and purification After stabilization of the anaerobic digestion process, i.e., after 2 months of operation under constant conditions, samples from all digesters were collected into sterile 15-mL Falcon tubes right before the daily addition of the substrate and used immediately for DNA extraction. Biomass was harvested by centrifugation of the samples at 20,000×g for 10 min, and the total DNA was extracted and purified by using a FastDNA® SPIN Kit for soil (MP Biomedicals, Germany). DNA quantity and purity were determined photometrically using a NanoDrop® ND-1000 UV/Vis spectral photometer (PeqLab, Germany) and by agarose gel electrophoresis. Cloning and sequencing of 16S rRNA genes Bacterial 16S rRNA gene fragments were amplified using the primers 27 F and 1492R (Lane 1991). PCR was carried out in 25.0 μL reaction mixtures containing 1.0 μL of 100-fold diluted template DNA (equivalent to 1.0–2.0 ng), 1.0 μL (5.0 pmol) of each primer, 1.0 μL of 99.5% dimethyl sulfoxide, 8.5 μL of nuclease free water, and 12.5 μL of Taq Master Mix (QIAGEN, Hilden, Germany). The cycle parameters were as follows: an initial denaturation at 94°C for 4 min, 30 cycles of 45 s at 94°C, 1 min at 58°C, 2 min at 72°C, and a final elongation at 72°C for 20 min. Archaeal 16S rRNA gene fragments were PCR-amplified using the forward primer UniArc8F (5’-YCY GKT TGATCC YGS CRG-3’) and the reverse primer UniArc931R (5’-CCC GCC AAT TCC TTT HAG-3’). Reaction mixtures were equivalent to the PCR for bacterial 16S rRNA genes, with the


exception that 1.0 μL of extra 50 mM MgCl2 was added. The cycle parameters were as described for the bacterial 16S rRNA genes but using an annealing temperature of 54°C. All primers were purchased from biomers.net (Ulm, Germany). PCR products were cleaned up with a QIAGEN PCR Purification Kit and cloned using a QIAGEN PCR Cloning Kit (QIAGEN, Hilden, Germany). For each bioreactor sample, 192 bacterial and 96 archaeal clones were picked up. Screening of positive clones by restriction analysis with HaeIII and hierarchical cluster analysis of restriction patterns were done as described previously (Kleinsteuber et al. 2006). For partial sequencing of bacterial clones, the 16S rRNA gene-specific primers 27F and 519R (Lane 1991) and vector-specific M13 primers were used. Archaeal clones were sequenced with vector-specific M13 primers. Sequencing was performed using the Big Dye Terminator Ready Reaction Cycle Sequencing Kit 1.1 (Applied Biosystems) on an ABI PRISM 3100 Genetic Analyzer. Assembling of single sequencing reads was accomplished with Sequencher 4.8 (Gene Codes Inc.). The BLASTN tool (www.ncbi.nlm.nih.gov/BLAST; Altschul et al. 1990) was used to search for similar sequences in the GenBank database, and the RDP Classifier (http:// rdp.cme.msu.edu; Wang et al. 2007) was used for taxonomic assignment. Phylogenetic analysis was accomplished with the ARB software, version 5.1 (www.arb-home.de; Ludwig et al. 2004). The determined sequences were initially aligned to the SILVA SSURef database Release 102 (www.arb-silva.de; Pruesse et al. 2007) by the ARB Positional Tree Server and added to the SILVA tree using the Quick Add Parsimony tool and applying specific filters for bacteria and archaea, respectively. The alignment was verified by comparison to the next relative sequences and corrected manually. The final position within the SILVA tree and bootstrap values were calculated by the Parsimony Interactive tool. The determined 16S rRNA gene sequences were deposited in the GenBank database under accession numbers HQ290266-290317. T-RFLP analysis Bacterial and archaeal 16S rRNA gene fragments were PCR-amplified as described above with the exception that the forward primers were labeled at the 5’-end with phosphoramidite fluorochrome-5-carboxyfluorescein. Amplicons were purified with the Wizard® SV PCR Clean-Up System (Promega, Mannheim, Germany) and quantified after electrophoresis in 1.5% agarose gels and ethidium bromide staining by the GeneTools program (Syngene, Cambridge, UK). The purified PCR products were then digested with the restriction endonucleases HaeIII or RsaI, respectively (New England Biolabs, Schwalbach, Germany), using 10 units of the respective


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enzyme for digesting 10 ng of archaeal and 20 ng of bacterial PCR product. Samples were incubated at 37°C over night and then precipitated with 0.1 volumes of 3 M sodium acetate (pH 5.5) and 2.5 volumes of absolute ethanol. Dried DNA samples were resuspended in 20 μL HiDi formamide containing 1.5% (v/v) GeneScan-500 ROX standard (Applied Biosystems). Samples were denatured at 95°C for 5 min and chilled on ice. The fragments were separated by capillary electrophoresis on an ABI PRISM 3100 Genetic Analyzer (Applied Biosystems). The lengths of the fluorescent terminal restriction fragments (T-RF) were determined using the GeneMapper V3.7 software (Applied Biosystems). Fluorescence signals of T-RF in the range of 50–800 bp were extracted, and peak areas were normalized according to (Abdo et al. 2006). Relative peak areas were determined by dividing the individual T-RF area by the total area of peaks within the range of 50 to 800 bp. Theoretical T-RF values of the representative phylotypes represented in the clone library were calculated using the NEB cutter (http://tools.neb.com/NEBcutter2) and confirmed experimentally by T-RFLP analysis using the corresponding clones as templates. Relative T-RF abundances of representative phylotypes were determined based on the relative peak areas of the corresponding T-RF.

Results Performance of the digesters Table 1 shows the operation parameters of the digesters containing DDGS as the substrate. The production of biogas stabilized after 2 months of operation under constant conditions with methane contents varying from 62.3% to 64.4%. The average biogas production, biogas composition, and other analytical parameters recorded during the last 3 weeks of operation are given in Table 2.

The addition of Fe3+ to the digesters for H2S precipitation caused a color change of the digesting mixture from yellow–brown to black due to the precipitation of insoluble iron sulfide. The supply of additional Fe3+ resulted in the removal of H2S from the gaseous phase, a higher biogas production and more stable methanogenesis reflected by lower VFA concentrations (Table 2). The causal connection becomes obvious when comparing the digesters nos. 6.2 and 6.6, which were identical in operation conditions except for the addition of FeCl3 to the latter. The average biogas production in digester no. 6.2 was 5.2 Lday−1 as opposed to 6.1 Lday−1 in digester no. 6.6. Furthermore, the lower biogas yield from digester no. 6.2 might also be a result of a shortage of micronutrients due to their precipitation in the presence of more H2S. Since some microorganisms inhabiting anaerobic bioreactors are facultative anaerobes, an aerobic pretreatment of the substrate with an inoculum from the appropriate digester was applied to see whether this resulted in more effective hydrolysis. Experiments with the bioreactors nos. 6.5 and 6.6 showed that such a pretreatment was inefficient or unnecessary, since the amount of biogas produced by both digesters did not vary significantly (5.9 vs. 6.1 Lday−1; Table 2). The effect of a higher OLR is seen when comparing the running parameters of digesters nos. 6.5 and 6.3 (Tables 1, 2). The higher OLR in no. 6.3 enhanced the biogas production from 5.9 to 13.7 Lday−1. The higher OLR results in the production of more organic acids. To avoid a rapid drop of pH in this digester, we used FerroSorp® DG as an additive precipitating H2S as non-toxic iron sulfide and stabilizing the pH because of its basic properties. Raw wheat stillage is separated into liquid and solid at an industrial scale to facilitate the drying process for DDGS production. Digester no. 6.4 was used to test whether the liquid phase can be used as the feedstock for biogas production. The OLR was identical to that of digester no.

Table 2 Process parameters measured during anaerobic digestion of DDGS Biogas composition pH Digester Biogas no. productiona (LN day−1) CH4 (%) CO2 (%) H2S (ppm)

Acid capacity Volatile fatty acids (mgL−1) NH4+-N −1 (gL ) (gL−1) Acetic acid Propionic acid Isobutyric acid

6.2 6.3 6.4

5.2±0.18 13.7±0.32 6.0±0.14

63.0±0.6 36.1±0.8 4513±71 64.4±0.6 35.1±0.4 34±4.9 62.7 ±0.5 36.6±0.7 264±15

7.51±0.07 0.94±0.07 7.68±0.04 1.25±0.11 7.17±0.03 0.57±0.05

149.5±8.1 70.1±7.2 0.3±0.1

64.3±4.4 2.7±0.12 0.2±0.1

1.7±0.02 n.d. n.d.

1.82±0.09 2.94±0.15 1.16±0.09

6.5 6.6

5.9±0.07 6.1±0.08

62.5±0.6 37.1±0.9 50±3.4 62.3±0.4 37.4±0.3 66±3.5

7.27±0.02 0.62±0.06 7.26±0.02 0.58±0.03

7.4±1.1 0.3±0.1

n.d. n.d.

n.d. n.d.

1.76±0.03 1.78±0.04

Average values of data points recorded during the last 3 weeks of operation are presented (n=21 for biogas production and composition; n=6 for the other parameters) n.d. not detectable a

Biogas production corrected for standard conditions (273.15 K and 101.325 kPa)

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6.5 (Table 1). Similar average biogas yields from both digesters (6.0 vs. 5.9 Lday−1; Table 2) indicated that raw wheat stillage is suited as feedstock for biogas production. Ammonium concentrations varied between 1.16 and 2.94 gL-1 and pH values between 7.17 and 7.68 in all tested digesters (Table 2). These values are characteristic of well-established anaerobic digestion systems (WPCF 1987; Gerardi 2003). In addition, low stationary concentrations of VFA confirmed that the anaerobic systems performed well, and VFA produced by acidogenic bacteria were almost completely funneled into methanogenesis, with the exception of the least efficient reactor no. 6.2 (Table 2). Bacterial diversity T-RFLP fingerprinting of 16S rRNA gene amplicons revealed high bacterial diversities in the five digesters, but strongly diverging species compositions. When regarding the T-RFs contributing at least 1% of the total T-RF peak area in at least one of the digesters, 43 and 42 operational taxonomic units (OTUs) were distinguished with the restriction enzymes HaeIII and RsaI, respectively. Figure 1 illustrates the results of the T-RFLP analysis with RsaI. Similar results were obtained with HaeIII (not shown). Cloned bacterial 16S rRNA gene amplicons were partially sequenced and their T-RF lengths determined in silico and confirmed experimentally. In total, 27 clones out of a library comprising 960 clones (192 clones from each digester) were selected for sequencing. These clones were divided into 13 OTUs based on their T-RFs and taxonomically assigned to the bacterial phyla Bacteroidetes, Actinobacteria, Firmicutes, Synergistetes, and Proteobacteria according to the RDP Classifier results (Table S1 in Supplemental Material). The

53 bp

100 90

Relative T-RF abundance (%)

Fig. 1 Phylogenetic diversity of bacteria in the five digesters according to relative T-RF abundances determined with the restriction enzyme RsaI. Only TRFs contributing at least 1% of the total T-RF peak area in at least one of the samples were included. Identification of T-RFs is based on partial sequences and experimental T-RF determination of cloned bacterial 16S rRNA genes (see Table S1 in Supplemental Material)

phylogenetic affiliations of the detected bacterial sequence types are shown in Figs. 2 and 3. As obvious from Fig. 1, many T-RFs could not be taxonomically assigned based on the clone library data, including some T-RFs with significant relative abundances. These points at a much higher bacterial diversity than reflected by the sequences compiled in Table S1. To assign more of the T-RF peaks, more clones need to be sequenced. Nevertheless, the most abundant and most diverse phylum in all digesters was the Bacteroidetes, with relative T-RF abundances of 32% in digester no. 6.2, 25% in no. 6.3, 41% in no. 6.4, 37% in no. 6.5, and 47% in no. 6.6. Within this phylum, five different OTUs, three of them assigned to the family Porphyromonadaceae, were detected in varying relative abundances (Fig. 1). The second most abundant phylum in digester no. 6.4, the Actinobacteria, comprised 19% of the total T-RF area. Three OTUs, one affiliated to the genus Brooklawnia and two to the genus Actinomyces, were detected within this group. In the other digesters, representatives of the Actinobacteria were less abundant with 1.7% relative T-RF abundance in digester no. 6.2, 4.4% in no. 6.3, less than 1% in no. 6.5, and 3.5% in no. 6.6. Two OTUs were assigned to the phylum Firmicutes— one to the family Peptococcaceae and one to another group of Clostridiales. The latter comprised 12% of the total T-RF peak area in digester no. 6.4, whereas in the other digesters the identified phylotypes affiliated to the Clostridia were less abundant (Fig. 1). The phylum Synergistetes was represented by one phylotype closely related to the genus Aminomonas. Its relative TRF abundance amounted to 9% in digester no. 6.6 and to 2% in no. 6.5, but it was not detected in the other digesters. Two

80 70 60 50 40 30 20 10 0 digester 6.2

digester 6.3

digester 6.4

digester 6.5

digester 6.6

55 bp

63 bp

Brooklawnia

82 bp

83 bp

97 bp

140 bp

144 bp

147 bp

168 bp

175 bp

180 bp

212 bp

Bacteroidetes I

Porphyromonadaceae II

314 bp

Porphyromonadaceae III

430 bp

443 bp

Synergistaceae

447 bp

448 bp

450 bp

452 bp

456 bp

Peptococcaceae

461 bp

464 bp

467 bp

468 bp

Bacteroidetes II

471 bp

Porphyromonadaceae I

Actinomyces sp. I

557 bp

563 bp

566 bp

636 bp

638 bp

Actinomyces sp. II

Clostridiales


Myxococcales 99 100 90

100

FJ416131, Bering Sea sediment clone HQ290305, clone bac6.5-2D6 100 FJ545582, Yellow Sea sediment clone AM040113, Wadden Sea sediment clone 100 AY225609, hydrothermal sediment clone EU488075, siliciclastic sediment clone 100 GU118797, coral clone AM882648, WWTP clone from oil-polluted water 90

Desulfurivibrio

100

66 AY005036, Desulfobulbus sp. oral clone CH031 DQ394978, harbor sediment clone VHS-B4-9 100 97 AM420151, subgingival plaque clone 402F09 EU844167, rumen fluid clone 100 90 HQ290317, clone bac6.6-1H12 HQ290311, clone bac6.6-2B9 76 FJ769451, anaerobic digester clone FP_C4 100 AY548789, Desulfobulbus propionicus X95180, Desulfobulbus elongatus 93

Deltaproteobacteria

Fig. 2 Phylogenetic tree of bacterial 16S rRNA gene sequences affiliated to the Deltaproteobacteria and Bacteroidetes. The determined partial 16S rRNA gene sequences (bold) were aligned to the SILVA SSURef database (release 102); the final position within the tree and bootstrap values were calculated using the ARB parsimony interactive tool. Only bootstrap values >50% are shown. Scale bar=10% nucleotide substitution

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Prolixibacter CU921945, municipal wastewater sludge clone HQ290312, clone bac6.6-2E6 CU922385, municipal wastewater sludge clone HQ290304, clone bac6.5-2D3 FJ516916, wetland sediment clone 100 FJ825441, biogas plant clone 100 HQ290300, clone bac6.4-2D3 AB175366, BSA-fed anaerobic digester clone BSA1B-12 74

100

100 100 100

Proteiniphilum

HQ290291, clone bac6.2-1E3 68 HQ290296, clone bac6.3-2B2 HQ290295, clone bac6.2-2F7 100 100 HQ290292, clone bac6.2-1F4 100 HQ290293, clone bac6.2-1G2 EU358739, pig manure biogas reactor clone 100 AM982599, pig saw dust spent bedding clone PISD-AJH04 FJ825516, biogas plant clone 75 DQ346489, uncultured compost bacterium clone 1B10 HQ290310, clone bac6.6-1B11 EF559147, mesophilic anaerobic digester clone EF586001, methanol-fed solid waste digester clone 96 CU919693, municipal watewater sludge clone 100 FJ205860, mesophilic biogas plant clone FJ825498, biogas plant clone 100 AY570690, Petrimonas sulfuriphila BN3 GU112187, pig manure biogas slurry clone 100 AM982594, pig saw dust spent bedding clone PISD-AIA06 98 HQ290306, clone bac6.5-2H8 94 HQ290298, clone bac6.3-2H11 HQ290303, clone bac6.4-2H6 66 AJ853536, landfill leachate clone GZKB42 AF349759, TCE-dehalogenating enrichment clone TCE5 98 AB234003, anaerobic reactor clone B-10 100 AB078842, Paludibacter propionicigenes

Bacteroidetes

100

0.10

sequence types affiliated to the Deltaproteobacteria were detected in the clone library generated from digester no. 6.6 but could not be quantified by T-RFLP profiles. One of these two OTUs was assigned to the genus Desulfobulbus, the other to the Desulfobacteraceae (Table S1). Archaeal diversity Archaea in the digesters were less diverse than bacteria as revealed by T-RFLP profiles. The T-RFs contributing at least 1% of the total T-RF peak area in at least one of the digesters

resulted in 11 and 13 OTUs with the restriction enzymes HaeIII and RsaI, respectively. Figure 4 shows the results obtained with HaeIII. Similar results were obtained with RsaI (not shown). Archaeal 16S rRNA gene clone libraries comprised 480 clones (96 from each sample), out of which 25 clones were selected for partial sequencing. According to these sequence data, nine OTUs were identified, eight of them affiliated to the phylum Euryarchaeota and one to the phylum Crenarchaeota (Table S2 in Supplemental Material). The phylogenetic affiliations of the euryarchaeote sequences are shown in


Fig. 3 Phylogenetic tree of bacterial 16S rRNA gene sequences affiliated to the Clostridia, Actinobacteria, and Synergistetes. The determined partial 16S rRNA gene sequences (bold) were aligned to the SILVA SSURef database (release 102); the final position within the tree and bootstrap values were calculated using the ARB parsimony interactive tool. Only bootstrap values >50% are shown. Scale bar=10% nucleotide substitution

Actinobacteria

AB274503, packed-bed reactor clone CFB-14 EU639200, thermophilic MFC clone SHBZ805 100 FN436047, thermophilic biogas reactor clone 97 AM947552, biowaste sludge clone CU918072, municipal wastewater sludge clone 100 HQ290307, clone bac6.6-1A3 95 HQ290309, clone bac6.6-1B2 100 AY297968, terephthalate-degrading sludge clone TTA_B12 AY661422, anaerobic hybrid reactor clone TTA_H151 88 AB091327, UASB granular sludge clone UT-1 99 AB091323, Pelotomaculum terephthalicicum AB232785, Pelotomaculum isophthalicicum 96 AB091324, UASB granular sludge clone JP 53 AY756143, biofilm clone 100 AB232815, propionate-degrading methanogenic consortium clone 100 EU266910, tar-oil contaminated aquifer sediment clone AB035723, Pelotomaculum thermopropionicum EU463221, naked mole-rat feces clone 100 AB088977, termite gut clone Rs-J39 100 EU469201, gazelle feces clone EU777322, okapi feces clone 95 AM696804, indoor dust clone AY850474, human intestine clone 99 EU467030, elephant feces clone DQ794601, human feces clone EU778533, river hog feces clone 100 EU775896, horse feces clone EU771525, baboon feces clone 54 EU465627, sheep feces clone 100 EU460021, colobus monkey feces clone EU469268, gazelle feces clone 97 98 FJ534954, waste activated sludge reactor clone 100 HQ290297, clone bac6.3-2H2 HQ290302, clone bac6.4-2E8 HQ290301, clone bac6.4-2E7 74 HQ290314, clone bac6.6-2E8 CU926578, municipal wastewater sludge clone 98 100 DQ196625, Brooklawnia cerclae BL-34 100 AJ003056, Propionimicrobium lymphophilum 100 HQ290294, clone bac6.2-2E10 99 HQ290308, clone bac6.6-1A9 57 AM084230, Actinomyces europaeus 100 FJ617539, Actinomyces sp. 7894GR 85 AJ249326, Actinomyces coleocanis DQ071541, Tetrao urogallus cecum clone 99 FJ393068, MFC anode enriched clone CU920445, municipal wastewater sludge clone 100 AY563469, wastewater MFC clone copi74 82 X80414, Actinomyces gerencseriae AF479270, Actinomyces israelii A1 100 EF558367, Actinomyces massiliensis 4401292 AJ697609, Actinomyces dentalis R18165 89 AJ575186, Actinomyces orihominis val1 100 DQ393001, reindeer rumen bacterium 6/9293-1 DQ072006, Actinomyces ruminicola 93 HQ290299, clone bac6.4-2C12 HQ290313, clone bac6.6-2H12

Clostridia

2046

Synergistes AF280829, bioreactor clone tbr4-7 CP001818, Thermanaerovibrio acidaminovor DSM6589 FN556061, Thermanaerovibrio sp. R101 DQ394907, harbor sediment clone EF559188, mesophilic anaerobic digester clone 100 EU887988, grass-silage fed leach bed reactor clone HQ290315, clone bac6.6-2H9 99 HQ290316, clone bac6.6-2H10 CU919853, municipal wastewater sludge clone AB175356, BSA-fed anaerobic digester clone BSA1B-02

90 100 100 100

Synergistetes

Aminiphilus

100

0.1

Figs. 5 and 6. The majority of clones belonged to the orders Methanomicrobiales and Methanosarcinales. No members of the orders Methanobacteriales and Methanococcales were detected. Hydrogenotrophic methanogens were predominantly represented by a phylotype affiliated to the genus Methanoculleus with relative T-RF abundances of 48% in digester no. 6.2 and 41% in no. 6.5. In the other digesters, this phylotype was less abundant with 9% in no. 6.3, 2% in no. 6.4, and 7% in no. 6.6. A second

representative of the hydrogenotrophic methanogens, Methanospirillum sp., was detected only in digester no. 6.6 in significant proportions (7%). Aceticlastic methanogens of the genus Methanosaeta dominated with 49% (no. 6.2), 79% (no. 6.4), 48% (no. 6.5), and 77% (no. 6.6) in all but one digester. An exception was digester no. 6.3 that was dominated by Methanosarcina sp. with a relative T-RF abundance of 84%. Members of the genus Methanomethylovorans were detected only in small proportions (1–4%) in digesters nos. 6.4, 6.5, and 6.6.

Appl Microbiol Biotechnol (2011) 89:2039–2052 100

Relative T-RF abundance (%)

Fig. 4 Phylogenetic diversity of archaea in the five digesters according to relative T-RF abundances determined with the restriction enzyme HaeIII. Only T-RFs contributing at least 1% of the total T-RF peak area in at least one of the samples were included. Identification of T-RFs is based on partial sequences and experimental T-RF determination of cloned archaeal 16S rRNA genes (see Table S2 in Supplemental Material)

2047 Methanomethylovorans

90

314 bp

80

290 bp

291 bp

Thermoprotei

70

Methanosarcina Methanomicrobiales

60

Methanosaeta II Methanosaeta I

50

Methanoculleus 53 bp

40 30 20 10 0 digester 6.2

Discussion The use of residual materials and biowaste as feedstock for biogas production is part of integrated concepts for biofuel production and biorefinery. While it is important to use high quality organic substrates for the production of certain biofuels, biogas can be generated from various municipal, industrial, agricultural, and food-processing waste products (Antoni et al. 2007). Here, we investigated the anaerobic digestion of DDGS, a by-product of bioethanol production, in lab-scale mesophilic reactors and analyzed the microbial community involved in the process. We tested the effectiveness of Fe3+ addition for H2S removal and aerobic pretreatment of distillers grains. The addition of Fe3+ to prevent H2S toxicity for microorganisms is a common operational practice (Gerardi 2003). With DDGS as a substrate, addition of Fe3+ as hydroxide (FerroSorp® DG) or chloride improved biogas yield and process stability. Moreover, the effect of such additives on the H2S content can facilitate biogas purification. Aerobic substrate pretreatment with appropriate inocula has been reported to be important for specific substrates (Angelidaki et al. 2002). However, for the anaerobic digestion of DDGS aerobic preincubation turned out to be dispensable. Bacteria that developed in the reactors during anaerobic digestion of DDGS were phylogenetically diverse, and no clear correlation with reactor parameters could be observed. However, generally the predominant phylotypes were assigned to the Bacteroidetes. Members of this phylum are commonly found in anaerobic waste sludge digesters (Chouari et al. 2005; Ariesyady et al. 2007; Rivière et al. 2009) and in biogas reactors fed with plant biomass (Krause et al. 2008; Kröber et al. 2009; Li et al. 2009), which is also illustrated by the phylogenetic tree (Fig. 2) showing sequences retrieved from anaerobic digesters that cluster together with the sequences from DDGS-fed digesters. The next cultured relatives of the Bacteroidetes

digester 6.3

digester 6.4

digester 6.5

digester 6.6

phylotypes detected in our digesters were the genera Proteiniphilum, Petrimonas, and Paludibacter within the family Porphyromonadaceae, and the sugar-fermenting, facultative anaerobe Prolixibacter bellariivorans (Holmes et al. 2007). Members of the Porphyromonadaceae produce various VFA from carbohydrates or proteins, e.g., the proteolytic bacterium Proteiniphilum acetatigenes (Chen and Dong 2005) produces acetate and propionate from various amino acids. Petrimonas sulfuriphila generates acetate during sugar fermentation but can also use elemental sulfur or nitrate as electron acceptor (Grabowski et al. 2005). Paludibacter propionicigenes produces propionate, acetate, and succinate during sugar fermentation (Ueki et al. 2006). In contrast to previous reports that detected Clostridia as the most prevalent bacterial class in full-scale mesophilic biogas plants fed with plant biomass (Klocke et al. 2007; Krause et al. 2008; Kröber et al. 2009), Clostridia were less abundant in our digesters. A clostridial phylotype related to sequences retrieved from various feces and activated sludge (Fig. 3) was abundant only in reactor no. 6.4. Members of the Clostridiales produce cellulosomes and are intensively involved in the degradation of recalcitrant microcrystalline cellulose, supporting acetogens and methanogens with the necessary short chain compounds (Lynd et al. 2002; Schwarz 2001). In our DDGS-fed digesters, cellulolytic Clostridia seemed to be of minor importance. Instead, another group of hydrolytic bacteria—members the predominantly aerobic or facultatively anaerobic Actinomycetales (Lynd et al. 2002)—were the second most abundant group. Actinomycetes are used in biological pretreatment of lignocellulosic material, enhancing the rates of delignification and hydrolysis (Taherzadeh and Karimi 2008). Actinomycetales as the predominant bacterial taxon were recently detected in lab-scale anaerobic digesters fed with food industry waste (Feng et al. 2010). As shown in Fig. 3, the three actinobacterial phylotypes identified in our digesters

2048 Fig. 5 Phylogenetic tree of archaeal 16S rRNA gene sequences affiliated to the Methanomicrobiales and Thermoplasmatales. The determined partial 16S rRNA gene sequences (bold) were aligned to the SILVA SSURef database (release 102); the final position within the tree and bootstrap values were calculated using the ARB parsimony interactive tool. Only bootstrap values >50% are shown. Scale bar=10% nucleotide substitution

Appl Microbiol Biotechnol (2011) 89:2039–2052 100 100

Methanosarcinales

AB248619, butyrate-degrading anaerobic reactor clone BHA09 AB175345, anaerobic BSA digester clone BSA1A-10 AB175351, anaerobic BSA digester clone BSA2A-06 HQ290290, clone arc6.6-F5 93 CU916380, municipal wastewater sludge clone 55 HQ290288, clone arc6.6-F1 100 EU358671, pig manure biogas digester clone AS21 99 EU358653, pig manure biogas digester clone AS03 EU358654, pig manure biogas digester clone AS04 56 90 AB494243, anaerobic digester sludge clone WA27 100 DQ785299, meromictic lake clone AB355094, Manzallah Lake water clone 99 AB233304, UASB granular sludge clone AB236087, anaerobic granular sludge clone MP-Eth-A EU888809, UASB reactor clone A05 AB233300, anaerobic digested sludge clone ND-2 54 AJ576220, landfill leachate clone GZK26 100 AB244307, fatty acids-degrading methanogenic consortium clon 100 AY196683, Methanospirillum hungatei 100 AB494258, anaerobic digester clone FA69 AB494256, anaerobic digester clone FA54 75 FJ205787, Bielefeld biogas plant clone A114 52 HQ290271, clone arc6.3-D6 AB175350, anaerobic BSA digester clone BSA2A-05 75 EU369626, landfill clone AY196674, Methanoculleus bourgensis CT573574, municipal WWTP clone Methano75 EU369613, landfill clone microbiales FJ205780, Bielefeld biogas plant clone A72 EF552185, solid waste anaerobic digester clone 75 FJ205766, Bielefeld biogas plant clone A32 HQ290270, clone arc6.2-H2 HQ290268, clone arc6.2-D9 HQ290266, clone arc6.2-A4 90 HQ290267, clone arc6.2-D1 CT573517, municipal WWTP clone HQ290269, clone arc6.2-D12 74 AB248609, butyrate-degrading anaerobic reactor clone BLA06 Y16382, Methanoculleus palmolei AF220343, oil reservoir enrichment clone E31B1 DQ787476, Methanoculleus receptaculi EU591667, methanogenic reactor clone MCSArc_A2 96 EU721748, petroleum reservoir clone 75 GU129124, acetate enrichment from oil bed, clone 29aM AB232790, propionate-degrading methanogeneic consortium clon AB233301, anaerobic digested sludge clone ND-4 EU591665, methanogenic reactor clone MSCArc_A1 100 CT573625, municipal WWTP clone HQ290279, clone arc6.4-F9 96 AB232800, propionate-degrading methanogenic consortium clone 96 M59134, Methanoculleus marisnigri 95 AF531178, Methanoculleus submarinus 97 AB038795, Methanoculleus chikugoensis 52 AB065297, Methanoculleus thermophilus AB288249, deep subsurface groundwater clone HC-E3 100 AB084243, solid waste landfill clone LCD CU917425, municipal wastewater sludge clone 100 EU093109, sheep rumen clone CSIRO-WA22 100 EU284787, reindeer rumen clone NRmetA8 99 EF055534, cattle rumen clone LGMJN45 82 EU284794, reindeer rumen clone NRmetB25 90 AY816987, pig manure storage pit clone DF53 92 HQ290274, clone arc6.3-E6 EU413603, reindeer rumen clone SRmetF5 Thermoplasmatatales 100 EF055533, cattle rumen clone LGMJN44 100 EF541184, cattle rumen clone 4 96 EU662692, manure pit sludge clone T19 52 AJ576235, landfill leachate clone GZK61 HQ290273, clone arc6.3-E2 0.10

were related to the anaerobic actinomycetes Actinomyces ruminicola, Actinomyces europaeus, and Brooklawnia cerclae. The latter is a member of the Propionibacteriaceae that forms propionate and acetate in the course of glucose fermentation (Bae et al. 2006). The obligate anaerobe A. ruminicola was isolated from cattle rumen and is able to hydrolyze xylan and starch, generating formate, acetate, and lactate as fermentation products (An et al. 2006). Particularly high proportions of Actinomyces, Brooklawnia, and the clostridial phylotype were detected in digester no. 6.4, which was set up with the liquid phase of the DDGS

suspension as substrate, indicating that these hydrolytic bacteria had been specifically enriched during breakdown of long-chain components of DDGS. We conclude that Actinobacteria, besides Bacteroidetes and Clostridia, play an important role in the hydrolysis and acidogenesis during anaerobic digestion of distillers grains. Another clostridial phylotype detected in minor proportions in the reactors nos. 6.2 and 6.3 was affiliated to the Peptococcaceae and closely related to members of the genus Pelotomaculum (Fig. 3). Some members of this genus, also referred to as Desulfotomaculum subcluster Ih (Imachi et al.

Appl Microbiol Biotechnol (2011) 89:2039–2052 Fig. 6 Phylogenetic tree of archaeal 16S rRNA gene sequences affiliated to the Methanosarcinales. The determined partial 16S rRNA gene sequences (bold) were aligned to the SILVA SSURef database (release 102); the final position within the tree and bootstrap values were calculated using the ARB parsimony interactive tool. Only bootstrap values >50% are shown. Scale bar=10% nucleotide substitution

2049 100

Methanomicrobiales

EF552176, solid waste digester clone HQ290272, clone arc6.3-B3 AY225109, manure biogas reactor clone M1.3 AJ831154, landfill sludge clone 210 90 AJ831071, landfill leachate clone 72 EF175701, landfill clone C13A EU857627, Nisargruna biogas plant clone HQ290275, clone arc6.3-H1 100 HQ290276, clone arc6.3-H2 EU358651, pig manure biogas digester clone AS01 AB494248, anaerobic digester sludge clone SA16 M59140, Methanosarcina thermophila 73 M59137, Methanosarcina acetivorans 87 AJ012094, Methanosarcina barkeri U20151, Methanosarcina mazei 100 AF432127, Methanosarcina lacustris AJ238648, Methanosarcina baltica 100 AJ012742, Methanosarcina semesiae HQ290277, clone arc6.4-B7 100 HQ290286, clone arc6.5-H2 100 HQ290285, clone arc6.5-G9 100 AB447876, anaerobic granular sludge clone AY526516, methanol-fed lab scale bioreactor clone M8 AY526519, methanol-fed lab scale bioreactor clone P10 AY672821, Methanomethylovorans thermophila 100 FJ898361, oilfield formation water clone ANB-170 100 GU129128, oil bed clone 47aM AF120163, Methanomethylovorans hollandica DMS1 77 100 AJ276437, Methanomethylovorans victoriae X51423, Methanosaeta concilii AB248605, butyrate-degrading anaerobic reactor clone BLA02 AB355101, Manzallah Lake water clone AB434766, methanogenic sludge clone AnDHS-A4 EF592687, high sulfate anaerobic bioreactor clone 2R2A13 EU721745, oil well clone 99 AB447772, anaerobic granular sludge clone 58 CU916345, municipal wastewater sludge clone HQ290282, clone arc6.5-B4 52 HQ290287, clone arc6.6-B10 100 HQ290283, clone arc6.5-F6 100 AJ576227, landfill leachate clone GZK39 95 AB244305, fatty acids-degrading methanogenic consortium clone HQ290278, clone arc6.4-F4 HQ290280, clone arc6.4-G1 HQ290281, clone arc6.4-G9 HQ290284, clone arc6.5-F10 CU916032, municipal wastewater sludge clone 100 AB266892, UASB sludge granules clone SwA12fl AB355127, Manzallah Lake sediment clone AF395423, acetae-enriched culture clone Arc No. 5 AY817738, Methanosaeta harundinacea 0.10

2006) are obligate syntrophs that oxidize propionate but only in co-culture with hydrogenotrophic methanogens or other hydrogen-consuming partners. Members of the genus Pelotomaculum play an important role in syntrophic propionate oxidation in anaerobic wastewater processes as well as in natural anaerobic environments (Narihiro and Sekiguchi 2007). A member of the recently recognized phylum Synergistetes (Jumas-Bilak et al. 2009) was mainly detected in digester no. 6.6. Synergistetes are widely distributed, albeit mostly in minor proportions, in various anaerobic ecosystems such as oral caves, gastrointestinal tracts, wastewater treatment

systems, soils, oil wells, and anaerobic sludge digesters (Chouari et al. 2005; Godon et al. 2005; Vartoukian et al. 2007; Hugenholtz et al. 2009). They have been identified as one of the bacterial core groups involved in anaerobic sludge digestion (Rivière et al. 2009). All described species within the Synergistetes are strict anaerobes fermenting amino acids (Jumas-Bilak et al. 2009; Hugenholtz et al. 2009). In our digesters, together with the aforementioned proteolytic members of the Porphyromonadaceae, these bacteria might utilize the peptide constituents of the distillers grains for acidogenesis.

2050

In summary, we can draw some conclusions on the key bacteria involved in the single stages of DDGS digestion: Hydrolysis and acidogenesis might be brought about mainly by several groups of the Bacteroidetes, together with Actinomycetes, Clostridiales, and Synergistetes, the latter mainly involved in acidogenesis. Furthermore, acetogenesis might be performed by syntrophic members of the Peptococcaceae and to a minor extent by sulfate-reducing bacteria affiliated to the genus Desulfobulbus (Fig. 2), which is known to oxidize organic acids incompletely to acetate (Widdel and Hansen 1992). Homoacetogens were not detected. However, due to the low coverage of the sequenced bacterial clones, not all sequence types represented by the various T-RF were identified and a substantial part of the bacterial diversity in the digesters remains undisclosed, even more as rare phylotypes might be missed by PCR-based detection methods. As expected, archaeal diversity was lower than bacterial diversity; and in contrast to the bacterial T-RFLP profiles, most archaeal T-RF could be assigned to phylotypes represented in the archaeal clone library (Fig. 4). The digesters nos. 6.4, 6.5, and 6.6 supplied with FeCl3 for sulfide precipitation exhibited higher archaeal diversity, whereas in digester no. 6.2 only two major methanogenic phylotypes developed—one affiliated to the hydrogenotrophic genus Methanoculleus and the other belonging to the aceticlastic genus Methanosaeta. Ammonium toxicity should not affect archaeal community composition since the NH4+-N concentrations in the effluents from bioreactors with the same OLR were similar. Also pH values were in the range supporting growth of methanogens and appropriate to avoid inhibitory effects of ammonium (Demirel and Scherer 2008). However, in digester no. 6.2 high H2S concentrations or accumulation of VFA might have inhibited other archaeal groups. Members of the genus Methanoculleus have been frequently detected as predominant methanogens in agricultural biogas plants (Krause et al. 2008; Kröber et al. 2009; Nettmann et al. 2010). As VFA concentrations were low in the reactors with higher archaeal diversity, we can surmise that syntrophic associations between acetogenic bacteria and aceticlastic archaea were better accommodated in the digesters nos. 6.4, 6.5, and 6.6. These syntrophic relationships led to immediate acetate utilization by distinct Methanosarcinales and to enhanced methane production. Additionally, the supply of trace elements together with Fe3+ additives might have improved the activity of methanogens. By contrast, in digester no. 6.2 acetic acid and propionic acid accumulated, indicating an inhibition of syntrophic propionate-oxidizers as well as inefficient utilization of acetate by methanogens, which is also reflected by a lower methane yield. Accordingly, hydrogenotrophic methanogens (but only represented by the genus Methanoculleus) had a higher


relative abundance, as a larger part of the acetate pool might be utilized via syntrophic acetate oxidation delivering hydrogen and carbon dioxide funneled into hydrogenotrophic methanogenesis (Karakashev et al. 2006). In the digesters nos. 6.4, 6.5, and 6.6, all functional groups of methanogens showed a higher diversity than in the less efficient reactor no. 6.2. Hydrogenotrophic methanogens were additionally represented by the genus Methanospirillum (especially in digester no. 6.6), whereas aceticlastic methanogens comprised two phylotypes affiliated to the genus Methanosaeta. Additionally, Methanosarcina was present in small proportions (Fig. 5). The higher relative abundance of Methanosaeta I in the well-performing digester no. 6.6 correlated with lower relative abundances of Methanosaeta II and Methanoculleus. A representative of the obligately methylotrophic genus Methanomethylovorans was also detected in minor proportions in digesters nos. 6.4, 6.5, and 6.6. Methanomethylovorans hollandica was firstly isolated from freshwater sediments with dimethyl sulfide as sole carbon and energy source and is also capable of growing on methanol, methylamines, and methanethiol (Lomans et al. 1999). Dimethyl sulfide and methanethiol originate from the degradation of sulfur-containing amino acids or from the methylation of sulfide during degradation of methoxylated aromatic compounds (Lomans et al. 2002). Thus, these organisms might be involved in methanogenesis from sulfur-containing biomass in our digesters. Other Euryarchaeota affiliated to the Thermoplasmatales (Fig. 5) and Crenarchaeota affiliated to the Thermoprotei (Table S2 in Supplemental Material) were also detected as minor constituents of the archaeal communities. As they are only distantly related to any cultured species it is difficult to draw conclusions on their ecophysiological roles. However, the phylotypes belonging to the Thermoplasmatales clustered together with environmental sequences retrieved from typical methanogenic habitats such as various rumen microbiota, manure, or landfill leachate (Fig. 5). Archaeal diversity in the iron hydroxide amended digester no. 6.3 was also low, but here the methanogenic subcommunity was dominated by a phylotype affiliated to the genus Methanosarcina. This result reflects an enrichment effect of aceticlastic archaea adapted to high acetate concentrations due to the high OLR. Methanosarcina spp. dominate in the presence of high acetate concentrations and are common in anaerobic digesters, whereas Methanosaeta spp. outcompete Methanosarcina spp. at slow turnover rates and low acetate concentrations due to their high affinity to acetate (Kendall and Boone 2006). The abundance of the Methanosarcina phylotype in digester no. 6.3 might also be related to its tolerance to higher ammonium and VFA concentrations (Karakashev et al. 2005; Demirel and Scherer 2008). Hence, the Methanosarcina sp. phylo-


type prevailing in digester no. 6.3 seems to be an indicator organism for an efficient and stable biogas process relying on high OLR and thus requiring high acetate turnover rates. Accordingly, at lower OLR the Methanosaeta I phylotype prevailing in digester no. 6.6 is characteristic of stable aceticlastic methanogenesis. In conclusion, our results show that distillers grains are a valuable feedstock for biogas production. Thus, the implementation of this biotechnology in the ethanol industry can improve the efficiency of biomass utilization, as the biogas derived from anaerobic digestion of distillers grains can be used for heating and generating electricity in ethanol producing plants. Acknowledgments This work was partially supported by a grant “Alğarış” from the Republic of Tatarstan (Russia) and a program “Development of Scientific Potential of High School” (project № 8159, Russia) to Ayrat Ziganshin. We gratefully acknowledge Ute Lohse for technical assistance and the helpful comments and suggestions of Jan Postel.

References Abdo Z, Schuette UM, Bent SJ, Williams CJ, Forney LJ, Joyce P (2006) Statistical methods for characterizing diversity of microbial communities by analysis of terminal restriction fragment length polymorphisms of 16S rRNA genes. Environ Microbiol 8:929–938 Ahn HK, Smith MC, Kondrad SL, White JW (2009) Evaluation of biogas production potential by dry anaerobic digestion of switchgrass–animal manure mixtures. Appl Biochem Biotechnol 160:965–975 Alper H, Stephanopoulos G (2009) Engineering for biofuels: exploiting innate microbial capacity or importing biosynthetic potential? Nat Rev Microbiol 7:715–723 Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ (1990) Basic local alignment search tool. J Mol Biol 215:403–410 An D, Cai S, Dong X (2006) Actinomyces ruminicola sp. nov., isolated from cattle rumen. Int J Syst Evol Microbiol 56:2043– 2048 Angelidaki I, Ahring BK, Deng H, Schmidt JE (2002) Anaerobic digestion of olive oil mill effluents together with swine manure in UASB reactors. Water Sci Technol 45:213–218 Antoni D, Zverlov VV, Schwarz WH (2007) Biofuels from microbes. Appl Microbiol Biotechnol 77:23–35 Ariesyady HD, Ito T, Okabe S (2007) Functional bacterial and archaeal community structures of major trophic groups in a fullscale anaerobic sludge digester. Water Res 41:1554–1568 Bae HS, Moe WM, Yan J, Tiago I, da Costa MS, Rainey FA (2006) Brooklawnia cerclae gen. nov., sp. nov., a propionate-forming bacterium isolated from chlorosolvent-contaminated groundwater. Int J Syst Evol Microbiol 56:1977–1983 Bagi Z, Ács N, Bálint B, Horváth L, Dobó K, Perei KR, Rákhely G, Kovács KL (2007) Biotechnological intensification of biogas production. Appl Microbiol Biotechnol 76:473–482 Bothast RJ, Schlicher MA (2005) Biotechnological processes for conversion of corn into ethanol. Appl Microbiol Biotechnol 67:19–25 Chen S, Dong X (2005) Proteiniphilum acetatigenes gen. nov., sp. nov., from a UASB reactor treating brewery wastewater. Int J Syst Evol Microbiol 55:2257–2261

2051 Chen Y, Cheng JJ, Creamer KS (2008) Inhibition of anaerobic digestion process: a review. Bioresour Technol 99:4044–4064 Chouari R, Le Paslier D, Daegelen P, Ginestet P, Weissenbach J, Sghir A (2005) Novel predominant archaeal and bacterial groups revealed by molecular analysis of an anaerobic sludge digester. Environ Microbiol 7:1104–1115 Cirne DG, Lehtomaki A, Bjornsson L, Blackall LL (2007) Hydrolysis and microbial community analyses in two-stage anaerobic digestion of energy crops. J Appl Microbiol 103:516–527 Demirel B, Scherer P (2008) The roles of acetotrophic and hydrogenotrophic methanogens during anaerobic conversion of biomass to methane: a review. Rev Environ Sci Biotechnol 7:173–190 Feng XM, Karlsson A, Svensson BH, Bertilsson S (2010) Impact of trace element addition on biogas production from food industrial waste— linking process to microbial communities. FEMS Microbiol Ecol 74:226–240 Franke-Whittle IH, Goberna M, Pfister V, Insam H (2009) Design and development of the ANAEROCHIP microarray for invesigation of methanogenic communities. J Microbiol Methods 79:279–288 Gerardi MH (2003) The microbiology of anaerobic digesters. Wiley, Hoboken Goberna M, Insam H, Franke-Whittle IH (2009) Effect of biowaste sludge maturation on the diversity of thermophilic bacteria and archaea in an anaerobic reactor. Appl Environ Microbiol 75:2566–2572 Goberna M, Gadermaier M, García C, Wett B, Insam H (2010) Adaptation of methanogenic communities to the cofermentation of cattle excreta and olive mill wastes at 37°C and 55°C. Appl Environ Microbiol 76:6564–6571 Godon JJ, Morinière J, Moletta M, Gaillac M, Bru V, Delgènes JP (2005) Rarity associated with specific ecological niches in the bacterial world: the ‘Synergistes’ example. Environ Microbiol 7:213–224 Grabowski A, Tindall BJ, Bardin V, Blanchet D, Jeanthon C (2005) Petrimonas sulfuriphila gen. nov., sp. nov., a mesophilic fermentative bacterium isolated from a biodegraded oil reservoir. Int J Syst Evol Microbiol 55:1113–1121 Holmes DE, Nevin KP, Woodard TL, Peacock AD, Lovley DR (2007) Prolixibacter bellariivorans gen. nov., sp. nov., a sugarfermenting, psychrotolerant anaerobe of the phylum Bacteroidetes, isolated from a marine-sediment fuel cell. Int J Syst Evol Microbiol 57:701–707 Hugenholtz P, Hooper SD, Kyrpides NC (2009) Focus: Synergistetes. Environ Microbiol 11:1327–1329 Imachi H, Sekiguchi Y, Kamagata Y, Loy A, Qiu Y-L, Hugenholtz P, Kimura N, Wagner M, Ohashi A, Harada H (2006) Non-sulfatereducing, syntrophic bacteria affiliated with Desulfotomaculum cluster I are widely distributed in methanogenic environments. Appl Environ Microbiol 72:2080–2091 Jumas-Bilak E, Roudiere L, Marchandin H (2009) Description of ‘Synergistetes' phyl. nov. and emended description of the phylum 'Deferribacteres' and of the family Syntrophomonadaceae, phylum 'Firmicutes'. Int J Syst Evol Microbiol 59:1028–1035 Karakashev D, Batstone DJ, Angelidaki I (2005) Influence of environmental conditions on methanogenic compositions in anaerobic biogas reactors. Appl Environ Microbiol 71:331–338 Karakashev D, Batstone DJ, Trably E, Angelidaki I (2006) Acetate oxidation is the dominant methanogenic pathway from acetate in the absence of Methanosaetaceae. Appl Environ Microbiol 72:5138–5141 Kendall MM, Boone DR (2006) The order Methanosarcinales. In: Dworkin M, Falkow S, Rosenberg E, Schleifer KH, Stackebrandt E (eds) The prokaryotes—a handbook on the biology of bacteria, 3rd edn. Springer, New York, pp 244–256 Kleinsteuber S, Riis V, Fetzer I, Harms H, Muller S (2006) Population dynamics within a microbial consortium during growth on diesel fuel in saline environments. Appl Environ Microbiol 72:3531– 3542

2052 Klocke M, Mahnert P, Mundt K, Souidi K, Linke B (2007) Microbial community analysis of a biogas-producing completely stirred tank reactor fed continuously with fodder beet silage as monosubstrate. Syst Appl Microbiol 30:139–151 Klocke M, Nettmann E, Bergmann I, Mundt K, Souidi K, Mumme J, Linke B (2008) Characterization of the methanogenic Archaea within two-phase biogas reactor systems operated with plant biomass. Syst Appl Microbiol 31:190–205 Krakat N, Westphal A, Schmidt S, Scherer P (2010) Anaerobic digestion of renewable biomass—thermophilic temperature governs population dynamics of methanogens. Appl Environ Microbiol 76:1842– 1850 Krause L, Diaz NN, Edwards RA, Gartemann KH, Krömeke H, Neuweger H, Pühler A, Runte KJ, Schlüter A, Stoye J, Szczepanowski R, Tauch A, Goesmann A (2008) Taxonomic composition and gene content of a methane-producing microbial community isolated from a biogas reactor. J Biotechnol 136:91–101 Kröber M, Bekel T, Diaz NN, Goesmann A, Jaenicke S, Krause L, Miller D, Runte KJ, Viehöver P, Pühler A, Schlüter A (2009) Phylogenetic characterization of a biogas plant microbial community integrating clone library 16S-rDNA sequences and metagenome sequence data obtained by 454-pyrosequencing. J Biotechnol 142:38–49 Lane DJ (1991) 16S/23S rRNA sequencing. In: Stackebrandt E, Goodfellow M (eds) Nucleic acid techniques in bacterial systematics. Wiley, Chichester, pp 177–203 Lee C, Kim J, Hwang K, O'Flaherty V, Hwang S (2009) Quantitative analysis of methanogenic community dynamics in three anaerobic batch digesters treating different wastewaters. Water Res 43:157–165 Li T, Mazéas L, Sghir A, Leblon G, Bouchez T (2009) Insights into networks of functional microbes catalysing methanization of cellulose under mesophilic conditions. Environ Microbiol 11:889–904 Liu FH, Wang SB, Zhang JS, Zhang J, Yan X, Zhou HK, Zhao GP, Zhou ZH (2009) The structure of the bacterial and archaeal community in a biogas digester as revealed by denaturing gradient gel electrophoresis and 16S rDNA sequencing analysis. J Appl Microbiol 106:952–966 Lomans BP, Maas R, Luderer R, Op den Camp HJM, Pol A, van der Drift C, Vogels GD (1999) Isolation and characterization of Methanomethylovorans hollandica gen. nov., sp. nov., isolated from freshwater sediment, a methylotrophic methanogen able to grow on dimethyl sulfide and methanethiol. Appl Environ Microbiol 65:3641–3650 Lomans BP, van der Drift C, Pol A, Op den Camp HJ (2002) Microbial cycling of volatile organic sulfur compounds. Cell Mol Life Sci 59:575–588 Lübken M, Gehring T, Wichern M (2007) Microbiological fermentation of lignocellulosic biomass: current state and prospects of mathematical modeling. Appl Microbiol Biotechnol 85:1643–1652 Ludwig W, Strunk O, Westram R, Richter L, Meier H, Yadhukumar BA, Lai T, Steppi S, Jobb G, Förster W, Brettske I, Gerber S, Ginhart AW, Gross O, Grumann S, Hermann S, Jost R, König A, Liss T, Lüßmann R, May M, Nonhoff B, Reichel B, Strehlow R, Stamatakis A, Stuckmann N, Vilbig A, Lenke M, Ludwig T, Bode A, Schleifer K-H (2004) ARB: a software environment for sequence data. Nucleic Acids Research 32:1363–1371 Lynd LR, Weimer PJ, van Zyl WH, Pretorius IS (2002) Microbial cellulose utilization: fundamentals and biotechnology. Microbiol Mol Biol Rev 66:506–577

Appl Microbiol Biotechnol (2011) 89:2039–2052 McKendry P (2002) Energy production from biomass (Part 1): overview of biomass. Bioresour Technol 83:37–46 Narihiro T, Sekiguchi Y (2007) Microbial communities in anaerobic digestion processes for waste and wastewater treatment: a microbiological update. Curr Opin Biotechnol 18:273–278 Nettmann E, Bergmann I, Mundt K, Linke B, Klocke M (2008) Archaea diversity within a commercial biogas plant utilizing herbal biomass determined by 16S rDNA and mcrA analysis. J Appl Microbiol 105:1835–1850 Nettmann E, Bergmann I, Pramschüfer S, Mundt K, Plogsties V, Herrmann C, Klocke M (2010) Polyphasic analyses of methanogenic archaeal communities in agricultural biogas plants. Appl Environ Microbiol 76:2540–2548 O'Sullivan CA, Burrell PC, Clarke WP, Blackall LL (2005) Structure of a cellulose degrading bacterial community during anaerobic digestion. Biotechnol Bioengin 92:871–878 Pruesse E, Quast C, Knittel K, Fuchs BM, Ludwig W, Peplies J, Glöckner FO (2007) SILVA: a comprehensive online resource for quality checked and aligned ribosomal RNA sequence data compatible with ARB. Nucleic Acids Res 35:7188–7196 Rivière D, Desvignes V, Pelletier E, Chaussonnerie S, Guermazi S, Weissenbach J, Li T, Camacho P, Sghir A (2009) Towards the definition of a core of microorganisms involved in anaerobic digestion of sludge. ISME J 3:700–714 Sawyer CN, McCarty PL, Parkin GF (1994) Chemistry for Environmental Engineering, 4th ed., McGraw-Hill, New York Schink B (1997) Energetics of syntrophic cooperation in methanogenic degradation. Microbiol Mol Biol Rev 61:262–280 Schwarz WH (2001) The cellulosome and cellulose degradation by anaerobic bacteria. Appl Microbiol Biotechnol 56:634–649 Taherzadeh MJ, Karimi K (2008) Pretreatment of lignocellulosic wastes to improve ethanol and biogas production: a review. Int J Mol Sci 9:1621–1651 Talbot G, Topp E, Palin MF, Masse DI (2008) Evaluation of molecular methods used for establishing the interactions and functions of microorganisms in anaerobic bioreactors. Water Res 42:513–537 Ueki A, Akasaka H, Suzuki D, Ueki K (2006) Paludibacter propionicigenes gen. nov., sp. nov., a novel strictly anaerobic, Gram-negative, propionate-producing bacterium isolated from plant residue in irrigated rice-field soil in Japan. Int J Syst Evol Microbiol 56:39–44 Vartoukian SR, Palmer RM, Wade WG (2007) The division “Synergistes”. Anaerobe 13:99–106 Wang Q, Garrity GM, Tiedje JM, Cole JR (2007) Naive Bayesian classifier for rapid assignment of rRNA sequences into the new bacterial taxonomy. Appl Environ Microbiol 73:5261–5267 Weiland P (2010) Biogas production: current state and perspectives. Appl Microbiol Biotechnol 85:849–860 Weiss A, Jérôme V, Burghardt D, Likke L, Peiffer S, Hofstetter EM, Gabler R, Freitag R (2009) Investigation of factors influencing biogas production in a large-scale thermophilic municipal biogas plant. Appl Microbiol Biotechnol 84:987–1001 Widdel F, Hansen TA (1992) The dissimilatory sulfate- and sulfurreducing bacteria, 2nd edn. Springer, New York WPCF (1987) Anaerobic sludge digestion. Manual of Practice No.19, 2nd edn. Water Pollution Control Federation, Alexandria, VA. Yang SJ, Kataeva I, Hamilton-Brehm SD, Engle NL, Tschaplinski TJ, Doeppke C, Davis M, Westpheling J, Adams MWW (2009) Efficient degradation of lignocellulosic plant biomass, without pretreatment, by the thermophilic anaerobe “Anaerocellum thermophilum” DSM 6725. Appl Environ Microbiol 75:4762–4769