Agricultural residues such as sugar beet pulp and citrus peel are rich in pectin, which contains galacturonic acid as a main monomer. Pectin-rich residues are underexploited as feedstocks for production of bulk chemicals or biofuels. The anaerobic, fermentative conversion of d-galacturonate in anaerobic chemostat enrichment cultures provides valuable information toward valorization of these pectin-rich feedstocks. Replicate anaerobic chemostat enrichments, with d-galacturonate as the sole limiting carbon source and inoculum from cow rumen content and rotting orange peels, yielded stable microbial communities, which were dominated by a novel Lachnospiraceae species, for which the name “Candidatus Galacturonibacter soehngenii” was proposed. Acetate was the dominant catabolic product, with formate and H2 as coproducts. The observed molar ratio of acetate and the combined amounts of H2 and formate deviated significantly from 1, which suggested that some of the hydrogen and CO2 formed during d-galacturonate fermentation was converted into acetate via the Wood-Ljungdahl acetogenesis pathway. Indeed, metagenomic analysis of the enrichment cultures indicated that the genome of “Candidatus G. soehngenii” encoded enzymes of the adapted Entner-Doudoroff pathway for d-galacturonate metabolism as well as enzymes of the Wood-Ljungdahl pathway. The simultaneous operation of these pathways may provide a selective advantage under d-galacturonate-limited conditions by enabling a higher specific ATP production rate and lower residual d-galacturonate concentration than would be possible with a strictly fermentative metabolism of this carbon and energy source.
IMPORTANCE This study on d-galacturonate metabolism by open, mixed-culture enrichments under anaerobic, d-galacturonate-limited chemostat conditions shows a stable and efficient fermentation of d-galacturonate into acetate as the dominant organic fermentation product. This fermentation stoichiometry and population analyses provide a valuable baseline for interpretation of the conversion of pectin-rich agricultural feedstocks by mixed microbial cultures. Moreover, the results of this study provide a reference for studies on the microbial metabolism of d-galacturonate under different cultivation regimes.
Conversion of agricultural and food-processing residues by open, mixed microbial communities is highly relevant for treatment of waste streams and increasingly also investigated as a strategy for production platform chemicals from low-value feedstocks. As an alternative to complete anaerobic conversion to methane and carbon dioxide, fermentative metabolism of these residues can yield more-valuable products, including organic acids, solvents, and transport fuels. Production of such compounds in nonaseptic open cultures has the potential to enable much lower capital and processing costs than currently achievable with pure-culture processes (1–3).
Continuously operated, nonaseptic cultures enrich for microbial consortia that thrive under the imposed conditions (1, 4). By carefully designing and optimizing reactor configurations and cultivation regimes, microbial populations that specifically and reproducibly yield desired product profiles can be enriched (5). Parameters such as culture pH, biomass retention time, and temperature, either in steady-state or dynamic cultivation regimes, are among the key parameters influencing population composition and product profiles (6, 7). An additional decisive parameter in determining product formation by enrichment cultures is feedstock composition. Cell wall polymers and storage carbohydrates represent the majority of the fermentable compounds in most plant-derived waste streams. Hydrolysis of these polymers by extracellular microbial enzymes generates monomers, which are the actual substrates for anaerobic fermentation. Model studies with enrichment cultures on individual monomeric substrates can generate valuable information on the metabolic pathways and product profiles that may be encountered under industrially relevant conditions.
Previous studies on anaerobic fermentation of plant biomass monomers by continuous anaerobic enrichment cultures focused mainly on hexose and pentose sugars released from the cellulose and hemicellulose fractions of plant biomass (8–10). However, depending on the feedstock, other monomers can represent a significant or even the major fraction of the fermentable compounds in biomass hydrolysates. In particular, sugar beet pulp, citrus peel, and apple pomace are rich in pectin, a polymer that contains d-galacturonic acid as a main constituent (11–14). Large-volume citrus peel waste streams originate from producing orange juice concentrates, which occurs mainly in the United States and Brazil. Sugar beet pulp and apple pomace are produced predominantly in Europe as waste streams of sugar beet and apple processing, respectively (15, 16). These pectin-rich agricultural residues are currently mostly dried and sold as cattle feed, but the cost of the drying process reduces profitability of this application. In 2016 approximately 35 million tons of citrus peel, 28 million tons of sugar beet pulp, and 2 million tons of apple pomace were produced in the world, which, by dry weight, contain 17%, 14%, and 16% galacturonate, respectively (12–14, 17). These large-volume residues represent a potential and currently underexploited feedstock for production of bulk chemicals or biofuels.
Intensively studied sugar substrates such as glucose, xylose, or arabinose can be fermented to commodity products, including ethanol and lactate, at near-theoretical yields. However, the currently documented biochemical pathways for galacturonate metabolism do not enable high carbon and electron yields of these fermentation products (18–20). A major constraint in known pathways for dissimilation of galacturonate, which is more oxidized than glucose, involves cleavage of a C6 intermediate into two C3 moieties, eventually leading to the redox-neutral generation of two moles of pyruvate from one mole of galacturonate (Fig. 1) (21–26). As a consequence, metabolic reactions beyond pyruvate need to be redox balanced, for example, by conversion of pyruvate into equimolar amounts of acetyl coenzyme A (acetyl-CoA) and formate (Fig. 1). Consistent with this observation, engineered microbial strains exhibit low carbon yields on galacturonate for compounds that are more reduced than pyruvate, such as ethanol and lactate (11, 19, 20, 27).
Despite its ecological and industrial significance as a plant-derived carbon source, fermentative conversion of galacturonate by anaerobic chemostat enrichment cultures has not been studied. The aim of this study was to characterize product profiles and microbial populations of open, mixed-culture chemostat cultures fed with d-galacturonate as the sole limiting substrate and compare these results to cultures fed with the much better studied neutral sugar glucose (8).
Enrichment of galacturonate-fermenting microbial cultures in anaerobic chemostats.
To study the composition and physiology of anaerobic microbial enrichment cultures on d-galacturonate, duplicate chemostat cultures were grown on a synthetic medium in which this pectin monomer was the sole carbon and energy substrate. Both bioreactors were started by adding anaerobic shake flask precultures on the same medium, which had been inoculated with rotting orange peel and cow rumen content. The bioreactors were operated as batch cultures until all galacturonate was consumed and then were switched to continuous operation at a dilution rate of 0.13 h−1. This dilution rate and other parameters were chosen to enable direct comparison with previous experiments in glucose-limited chemostat cultures (8). Subsequently, product concentrations and carbon dioxide production were monitored over a period of 6 weeks. Constant rates of CO2 production and of metabolite concentrations in the chemostats indicated that a “metabolic steady state” was reached after 2 weeks of continuous cultivation (approximately 63 generations of the overall microbial population) (Fig. 2). In the steady-state cultures, the d-galacturonate concentration remained below the detection limit of 0.1 mM, consistent with the medium composition, which was designed to make d-galacturonate the growth-limiting nutrient. After 1 month of steady-state operation (approximately 135 generations), five samples were taken over a period of 5 days for further physiological and metagenomic analysis.
Product profiles in chemostat enrichment cultures suggest the involvement of acetogenesis in anaerobic galacturonate metabolism.
To investigate galacturonate metabolism, fermentation products were analyzed in the culture supernatants and off-gas of the steady-state enrichment cultures. In both reactors, acetate was the main catabolic product of d-galacturonate fermentation, with additional production of H2 and formate (Table 1). Average recoveries of carbon (92%) and electrons (95%) in the steady-state cultures indicated that the major fermentation products were accounted for (see Table S1 in the supplemental material).
TABLE 1 Comparison of product yields of anaerobic chemostat enrichment cultures on galacturonate and glucosea
Replicate enrichment cultures (bioreactors 1 and 2) were grown on synthetic medium with galacturonate as the limiting substrate, pH 8, at 30°C and at a dilution rate of 0.13 h−1.
Yields of biomass and products on galacturonate or glucose are expressed as Cmol Cmol−1 unless stated otherwise. Data represent averages ± SD for five sequential measurements during steady state.
Data for glucose-limited cultures (grown under conditions identical to those of glucuronate-limited enrichment) are from Temudo et al. (8).
The sum of the acetyl-CoA derivatives acetate, butyrate, and ethanol in moles of acetyl-CoA per Cmol of substrate.
In contrast to previously analyzed anaerobic chemostat enrichment experiments on glucose, which were grown under the same conditions (8) as used in the present study, the galacturonate-grown enrichment cultures did not produce detectable amounts of reduced fermentation products such as butyrate or ethanol (Table 1). Qualitatively, the formation of acetate, formate, CO2, and H2 as major fermentation products was consistent with previously described pathways for bacterial galacturonate metabolism (Fig. 1). However, in contrast to the product distribution expected from such pathways and to previous observations on glucose-grown chemostat enrichment cultures (8), the sum of the acetyl-CoA-derived products significantly differed from the combined formate and H2 concentrations. In bioreactors 1 and 2, the combined yields of hydrogen and formate on d-galacturonate (mole mole−1) were 37% ± 2% and 37% ± 1% lower, respectively, than the corresponding yields of acetate (Table 1). The observed metabolite profiles would be consistent with a contribution of acetogenesis via the Wood-Ljungdahl (WL) pathway, which enables conversion of fermentatively produced hydrogen (or alternatively, reduced ferredoxin derived from a pyruvate:ferredoxin oxidoreductase, EC 126.96.36.199) and carbon dioxide to acetate (equation 1). The combination of acetogenesis with fermentative conversion of d-galacturonate into acetate via a classical modified Entner-Doudoroff (ED) route (equation 2; Fig. 1) allows for conversion of d-galacturonate to acetate without net production of hydrogen (equation 3; Fig. 3).
Based on the assumption that d-galacturonate catabolism in the enrichment cultures indeed reflected simultaneous d-galacturonate fermentation via an ED pathway and acetogenesis via the WL pathway, the distribution of carbon and electrons over both pathways can be easily estimated from measured rates of d-galacturonate consumption and of hydrogen, formate, and acetate production in the steady-state cultures (three equations with two unknowns ). Based on these calculations (see calculations in the supplemental material), the average specific rates of acetate formation from pyruvate generated in the ED pathway (qacetate,ED) and of acetogenesis via the WL pathway (qacetate,WL) were estimated to be 6.9 ± 0.4 mmol (g biomass)−1 h−1 and 1.7 ± 0.2 mmol (g biomass)−1 h−1, respectively. In such a scenario, 16% ± 3% of the produced hydrogen and formate generated from d-galacturonate would have been used for acetogenesis (Fig. 3).
Population analysis of anaerobic d-galacturonate-fermenting enrichment cultures.
To study microbial population dynamics during enrichment and to assess if the hypothesis outlined above was supported by the presence of acetogenic microorganisms, 16S-rRNA gene amplicon sequencing was performed on samples from both bioreactors. The inoculum used to inoculate both bioreactors contained species related to the genera Clostridiumsensu stricto and Lachnoclostridium as well as to uncultured Lachnospiraceae (Fig. 4). After complete consumption of d-galacturonate in the batch phase preceding the continuous culture, bioreactor 1 was dominated by bacteria related to Raoultella and Yersinia, with the latter showing a 99% identity to Yersinia massiliensis in the SILVA small subunit (SSU) database release 128. At the same sampling time, bioreactor 2 was dominated by bacteria related to Klebsiella and Enterobacter. Despite this difference in population composition, product profiles were similar for the two bioreactors at this time point, with acetate as the dominant organic catabolic product and formate as the second major product (Fig. 2).
Although the microbial community compositions were different at the start of the continuous enrichment, similar populations developed during prolonged continuous operation of the two bioreactors. In both reactors, a bacterium related to Lachnotalea became dominant, with a side population of a Klebsiella-related bacterium (Fig. 4). The steady-state communities in the two replicates were highly similar, indicating that the chosen cultivation conditions generated a selective advantage for these species and their conversions.
Community distribution determination by FISH analysis.
To further quantify the abundance of dominant community members during the steady-state measurements, fluorescence in situ hybridization (FISH) analysis (29, 30) was performed (Fig. 5). A general probe (EUB338) was used to stain all bacterial cells, while based on the results of the 16S rRNA gene analysis, a probe specific for the Lachnospiraceae family (Lac435) and a probe for the Enterobacteriaceae family (ENT) were used to quantify bacteria related to Lachnotalea and Klebsiella, respectively. Microorganisms belonging to the Lachnospiraceae were found to be abundant in both reactors, with another abundant population of bacteria belonging to the Enterobacteriaceae (Fig. 4 and 5). These observations correlated with the identification of a Lachnotalea- and a Klebsiella-related bacterium, respectively, in the 16S rRNA gene analysis.
Metagenomic analysis of pathways and species in the enrichment cultures.
To investigate whether the enriched Lachnotalea-related bacterium harbored genes encoding enzymes of the WL pathway and for DNA-based identification of the dominant microorganisms, a metagenomic analysis was performed on the steady-state cultures. DNA of the entire community was extracted from both steady-state enrichment cultures and sequenced. In total, 14.9 million paired-end reads remained posttrimming, and 7.9 Gbp of sequenced bases were obtained for each of the enrichments. After de novo assembly and metagenome binning-based sequence composition and differential coverage, draft genomes of the dominant microorganisms were assembled (31–35), checked for completeness and contamination, and annotated with the RAST server (36–39).
The genome with the highest coverage (577-fold ± 94-fold; completeness, 98%) in both enrichments was identified to be a member of the Lachnospiraceae by analysis of the small subunit rRNA gene of the constructed genome. An identity of 93% was found within the SILVA SSU database release 132 with the closest cultured relative, Lachnotalea glycerini, indicating that the dominant microorganism belongs to a novel genus within the Lachnospiraceae family (40, 41). We propose the name “Candidatus Galacturonibacter soehngenii” (Ga′.lac.tu.ro.ni.bac.ter. N.L. masc. n., galacturoni, referring to the utilization of galacturonate; bacter, the equivalent of Gr. neut. n. bactrum, a rod or staff, referring to the shape; soeh.ng.en′.ii., N.L. masc. n. gen., named after Nicolaas Söhngen, first Ph.D. student in Delft University of Technology and pioneer in anaerobic microbiology) for this novel microbe. The assembled and annotated genome of this organism harbored all structural genes for the enzymes of the canonic ED pathways for d-galacturonate catabolism, as well as multiple signature genes encoding enzymes of the WL pathway for acetogenesis, including fhs, which encodes formate-tetrahydrofolate ligase (EC 188.8.131.52) and is specific for the WL pathway (42–44) (Table 2). However, as shown in Table 2, the genes cooS, encoding carbon-monoxide dehydrogenase (EC 184.108.40.206), and acsBCD, which encodes the CO-methylating acetyl-CoA synthase complex (EC 220.127.116.11), were not identified.
TABLE 2 Genesa of the Wood-Ljungdahl and adapted Entner-Doudoroff pathways identified in the genome of “Candidatus G. soehngenii”
Genes with the highest annotation found and their corresponding expected value (E value) are listed.
—, no homologues were identified.
A complete set of WL pathway genes were identified in the metagenomic data set within the genome of a species closely related to a Sporomusa species (99% identity found in SILVA SSU database release 132), within a genus known for harboring homoacetogenic bacteria (45). However, the genome of the Sporomusa species had a coverage of only 9-fold ± 2-fold and a completeness of 80%. This coverage corresponds to approximately 1% ± 0% of the total coverage. The other genome with a high coverage (166-fold ± 12-fold; completeness, 100%), was closely related to Klebsiella oxytoca (100% identity of the 16S rRNA gene in the SILVA SSU database release 132). This observation was consistent with the 16S rRNA gene amplicon sequencing data, which showed a side population of a bacterium closely related to Klebsiella species. The K. oxytoca genome did not harbor any WL pathway genes but, in accordance with literature (46), did contain all the genes for the adapted ED pathway for d-galacturonate metabolism.
The identification of acetate as the dominant catabolic product of anaerobic chemostat enrichment cultures on d-galacturonate (Table 1) is consistent with early studies on the metabolism of pectin-fermenting bacteria (47–49). The narrow product range of the currently described d-galacturonate-fermenting organisms is likely to be a consequence of the redox-neutral conversion of d-galacturonate to pyruvate via a modified Entner-Doudoroff (ED) pathway (Fig. 1) (18). In contrast, glucose fermentation, in which oxidative glycolytic pathways generate reduced cofactors that are subsequently reoxidized by reduction of pyruvate and/or acetylphosphate, generates a wide diversity of fermentation products. In enrichment cultures on glucose, performed under conditions identical to those used in the present study, a 1:2 molar ratio of butyrate to formate or hydrogen in such cultures was consistent with a classical Embden-Meyerhof glycolysis and subsequent reduction of pyruvate (Table 1) (8, 50).
When d-galacturonate is metabolized to pyruvate via a modified ED pathway, subsequent redox-neutral conversion of pyruvate into acetate and formate can occur via pyruvate formate-lyase (EC 18.104.22.168), phosphate acetyltransferase (EC 22.214.171.124), and acetate kinase (EC 126.96.36.199) (C6H10O7 → 2 C2H4O2 + 2 HCOOH), with a potential further conversion of formate to hydrogen and CO2 by formate hydrogen-lyase (EC 188.8.131.52 and EC 184.108.40.206) (Fig. 1) (26, 50, 51). However, since the 1:1 molar ratio of acetate to formate or hydrogen expected in such a scenario was not observed, we hypothesized that, in the chemostat enrichment cultures, acetogenesis via the Wood-Ljungdahl (WL) pathway converted some of the hydrogen and CO2 produced during conversion of pyruvate to acetate.
The enrichment cultures contained different species suspected to be capable of acetogenesis. Based on a sequence coverage estimate, a member of the Sporomusa genus, which is well known to harbor autotrophic acetogens (45), made up less than 1% of the population in the enrichment cultures. If this bacterium was solely responsible for the inferred rates of acetogenesis in the cultures, its specific activity would have to exceed previously reported rates of acetogenesis (52, 53) by at least 1 order of magnitude. This makes it likely that another bacterium was responsible for the majority of the acetogenesis.
The dominance of “Candidatus G. soehngenii,” which was clearly shown in both the 16S rRNA gene (Fig. 4) and metagenome analysis, was not matched by the FISH analysis (Fig. 5). This discrepancy was most probably due to the reported difficulty of staining Gram-positive microorganisms (54, 55). We therefore assume that “Candidatus G. soehngenii,” while underrepresented in the FISH analysis, was indeed dominant and responsible for the main conversions observed in both bioreactors.
After the onset of continuous cultivation, “Candidatus G. soehngenii” gradually replaced other microorganisms (Fig. 3). The genome of this organism, which became dominant at the end of duplicate chemostat enrichment experiments, harbored most genes of the WL pathway. However, we did not identify genes with a high sequence similarity to known cooS and acsBCD genes respectively encoding carbon-monoxide dehydrogenase (EC 220.127.116.11) and CO-methylating acetyl-CoA synthase complex subunits (EC 18.104.22.168) (42). Genomes of Lachnospiraceae species remain underexplored and contain many nonannotated genes, and the possibility of finding novel functional homologues of the Wood-Ljungdahl pathway might be interesting for future research (56). Alternatively, the “missing” cooS and acsBCD genes may be an artifact of the metagenome assembly and analysis. All traditional methods were used in an attempt to isolate “Candidatus G. soehngenii” (for a detailed list of isolation methods, see Table S2 in the supplemental material), but none was successful. We postulate that “Candidatus G. soehngenii” is very well adapted to growth at limiting substrate concentrations (K-strategist) as occurring in a chemostat-operated bioreactor (57). Alternative isolation procedures not relying on batch growth might be needed for successful isolation. Although confirmation will require isolation of a pure culture, we expect “Candidatus G. soehngenii” to be capable of simultaneous fermentation of d-galacturonate and acetogenesis.
Microbial competition is often interpreted in terms of the affinity for a single growth-limiting nutrient (qsmax/Ks) (58). Under nutrient-limited conditions, simultaneous utilization of mixed substrates enables growth at lower concentrations of each of the substrates than would be possible when growth is limited by a single substrate (59, 60). In this way, reconsumption of hydrogen could enable lower residual galacturonate concentrations in the chemostat cultures, thereby conferring a selective advantage to “Candidatus G. soehngenii.” Additionally, the net ATP yield of d-galacturonate fermentation via an ED-type pathway amounts to 3 mol ATP (mol galacturonate)−1 or 1.5 mol ATP (mol acetate)−1, while the supplemental formation of acetate via the WL pathway is estimated to yield an additional 0.33 mol ATP (mol acetate)−1, assuming chemiosmotic energy conservation due to the generation of a transmembrane electrochemical ion gradient (61). With a constant growth rate under chemostat operations, the total ATP flux, qATP, required for growth will also remain constant. When the ATP is supplied by both the adapted ED pathway and the WL pathway, the relative qATP derived from the ED pathway can decrease, without lowering the overall qATP. Lowering the flux through the ED pathway decreases the galacturonate biomass specific uptake rate (qs), and in accordance to the substrate uptake kinetics, shown in equation 4, this will decrease the galacturonate concentration (Cs) in the bioreactor (62).
The very slow replacement of nonacetogenic bacteria in the chemostat enrichment cultures is in line with the relatively small impact of acetogenesis on the kinetics and energetics of d-galacturonate metabolism. Moreover, the dilution rate of the chemostat cultures was at the upper end of specific growth rates reported for typical heterotrophic homoacetogens on (semi-) defined media (63–65). Additionally, sparging of the reactors with nitrogen gas may have stripped H2 formed by d-galacturonate fermentation. Estimated hydrogen partial pressures in the reactors were 2 × 10−3 to 5 × 10−3 atm, resulting in a Gibbs free energy change of −17.3 ± 24.0 kJ mol−1 for the WL pathway (for calculations, see the supplemental material) (66, 67). With the Gibbs free energy change close to the minimal driving force for catabolic reactions, this low in situ hydrogen partial pressure may have limited the rate of H2 oxidation in acetogenesis. This interpretation is consistent with the dominance of fast-growing nonacetogenic bacteria (e.g., Klebsiella and Clostridium species) during the batch phase that preceded the chemostat enrichment.
Although involvement of the WL pathway has been demonstrated in anaerobic chemostat enrichment cultures on glucose at a low dilution rate (0.05 h−1) (66, 68), such an involvement was not observed at higher dilution rates (under operational conditions similar to those used in this study ). This observation suggests that potentially lowering the residual glucose concentration by simultaneous acetogenesis did not provide a selective advantage in anaerobic enrichment cultures fed with glucose. More research is required to elucidate this difference in usage of the Wood-Ljungdahl pathway in glucose- and galacturonate-limited cultures. Although “Candidatus G. soehngenii,” which became dominant in the d-galacturonate-limited enrichment cultures in this study, differed from the Clostridium quinii that dominated similar cultures grown on glucose as the carbon source (69), both are members of the Clostridiales. This observation illustrates the importance of these organisms in fermentative conversion of carbohydrates and related compounds in carbon-limited, anaerobic environments (69).
MATERIALS AND METHODS
A continuously stirred reactor of 1.2-liter capacity (0.5-liter working volume) was used (mechanical stirring, 300 rpm). Water was recirculated to maintain a constant temperature at 30°C. The reactor was sparged with nitrogen gas at a flow rate of 120 ml min−1 to maintain anaerobic conditions. The pH was controlled at pH 8 ± 0.1 by automatic titration (ADI 1030 Bio controller) with a 1 M NaOH solution. The dilution rate was 0.13 h−1, and the working volume was kept constant by peristaltic effluent pumps (Masterflex; Cole-Parmer, Vernon Hills, IL, USA) coupled to electrical level sensors.
To characterize the product spectrum, the reactor was run in continuous mode until stable product composition and biomass concentration were established. Steady state was reached after 2 weeks (63 generations). On-line analysis of system stability was achieved by online monitoring of the gas productivities, in the form of CO2, and base addition. On-line gas detection of CO2 was done with a Rosemount Analytical NGA 2000 MLT 1 multicomponent analyzer (infrared detector). Data acquisition was achieved with SCADA software (Sartorius BBI systems MFCS/win 2.1). When these rate measurements were stable or varied within a limited range (±10%) without a trend showing increase or decrease, a set of samples were taken during the subsequent cycles. The concentrations of soluble organic fermentation products, hydrogen produced, and biomass in the reactor volume were determined.
The reactor was inoculated (1%, vol/vol) with open mixed precultures started with a mixture of two different sources. The first inoculum was obtained from the content of rumen offal of a grass-fed cow (Est, Netherlands). The second inoculum was a sample from organic waste containing a large amount of citrus peels (Orgaworld Nederland B.V., Netherlands). Shake flasks were inoculated in duplicate with compost (2%, vol/vol) and rumen extract (2% vol/vol), and the initial pH was adjusted to 8 with a 2 M KOH solution. The shake flasks were cultivated in an anaerobic chamber (Bactron III; Shel Lab, Cornelius, USA) at 30°C with a gas composition of 89% N2, 6% CO2, and 5% H2.
The cultivation medium contained the following (in grams liter−1): d-galacturonate, 4.0; NH4Cl, 1.34; KH2PO4, 0.78; Na2SO4·10H2O, 0.130; MgCl2·6H2O, 0.120; FeSO4·7H2O, 0.0031; CaCl2, 0.0006; H3BO4, 0.0001; Na2MoO4·2H2O, 0.0001; ZnSO4·7H2O, 0.0032; CoCl2·H2O, 0.0006; CuCl2·2H2O, 0.0022; MnCl2·4H2O, 0.0025; NiCl2·6H2O, 0.0005; EDTA, 0.20. The d-galacturonate and mineral solutions were prepared and fed separately. The d-galacturonate solution was sterilized by filtration (0.2 μm Mediakap Plus; Spectrum Laboratories, Rancho Dominguez, CA, USA), and the mineral solution was autoclaved for 20 min at 121°C. Three milliliters PluronicPE 6100 antifoam (BASF, Ludwigshafen, Germany) was added per 40 liters mineral solution to avoid excessive foaming.
Analytical methods of substrate and extracellular metabolites.
To determine substrate and extracellular metabolite concentrations, reactor sample supernatant was obtained by centrifugation of culture samples (Biofuge Pico; Heraeus, Hanau, Germany) and subsequent filtration (0.2 μm Millex-HV; Millipore-Merck, Darmstadt, Germany). Concentrations of galacturonate and extracellular metabolites were analyzed using an Agilent 1100 Affinity high-pressure liquid chromatography (HPLC) machine (Agilent Technologies, Amstelveen, Netherlands) with an Aminex HPX-87H ion-exchange column (Bio-Rad, Hercules, CA, USA) operated at 60°C with a mobile phase of 5 mM H2SO4 and a flow rate of 0.6 ml min−1. Measurements of H2 and CO2 during steady-state analysis were performed off-line using gas bags (3.8 liters; Tedlar, Saint Gobain, France) and analyzed using a mass spectrometer (Prima BT MS; Thermo Scientific, USA).
Culture dry weight was measured by filtering 20 ml of culture broth over predried and preweighed membrane filters (0.2-μm Supor-200; Pall corporation, New York, NY, USA), which were then washed with demineralized water, dried in a microwave oven (Easytronic M591; Whirlpool, Benton Harbor, MI, USA), and weighed again. Carbon balances and electron balances were established based on the number of carbon atoms and electrons per mole, while a standard biomass composition of CH1.8O0.5N0.2 was assumed (70).
Microbial community stability analysis.
The bacterial DNA for the community analysis over time was obtained by extracting 2 ml broth and pelleting the biomass before storing it at −80°C for further analysis. The DNA was extracted using the UltraClean DNA isolation kit (MOBIO laboratories Inc., USA) as per the manufacturer's instructions. The 16S rRNA genes of days 1 and 38 of bioreactor 1 and day 38 of bioreactor 2 were analyzed using amplicon sequencing with an Illumina HiSeq (Novogene Bioinformatics Technology Co., Ltd.); for this, the primers 341F (5′-CCTAYGGGRBGCASCAG-3′) and 805R (5′-GGACTACNNGGGTATCTAAT-3′) were used, generating 250-bp paired-end reads. Sequences were analyzed according to the method of van den Berg et al. (71). Representative sequences for each operational taxonomic unit were submitted for BLAST analysis under default conditions. The extracted genomic DNAs of the inoculum, from days 9 and 14 of bioreactor 1 and days 1, 9, and 14 of bioreactor 2, were subsequently used for a two-step PCR targeting the 16S rRNA gene of most bacteria and archaea. For this, the primers U515F (5′-GTGYCAGCMGCCGCGGTA-3′) and U1071R (5′-GARCTGRCGRCRRCCATGCA-3′) were used according to the method of Wang et al. (72). The first amplification was performed to enrich for 16S rRNA genes, using a quantitative PCR assay with 2× iQ SYBR green Supermix (Bio-Rad, CA, USA), 500 nM primers each, and 1 to 50 ng of genomic DNA was added per well to a final volume of 20 μl. The thermocycles consisted of a first denaturation at 95°C for 5 min, 20 cycles at 95°C for 30 s, 50°C for 40 s, and 72°C for 40 s, and a final extension of 72°C for 7 min. 454-adapters (Roche) and MID tags at the U515F primer were added to the produced PCR products. The second amplification was similar to the first one, with the exception of the use of Taq PCR master mix (Qiagen Inc., CA, USA); the program was run for 15 cycles, and the template (the product from step one) was diluted 10 times. After the second amplification, the PCR products were pooled equimolar and purified over an agarose gel using a GeneJET gel extraction kit (Thermo Fisher Scientific, Netherlands). The resulting library was sent for 454 sequencing and run in 1/8 lane with titanium chemistry by Macrogen Inc. (Seoul, South Korea). After analysis, the reads library was imported into CLC genomics workbench v7.5.1 (CLC Bio, Aarhus, DK), quality trimmed (limit, 0.05) to a minimum of 200 bp and an average of 284 bp, and demultiplexed. A build-it SILVA 123.1 SSURef Nr99 taxonomic database was used for BLASTn analysis on the reads under default conditions. The top result was imported into an Excel spreadsheet and used to determine taxonomic affiliation and species abundance.
Fluorescence in situ hybridization.
FISH was performed as described in reference 73, using a hybridization buffer containing 25% (vol/vol) formamide. Probes were synthesized and 5′ labeled either with the Fluos dye or with one of the sulfoindocyanine dyes Cy3 and Cy5 (Thermo Hybaid Interaciva, Ulm, Germany) (Table 3). The general probe EUB338 labeled with Cy5 was used to indicate all eubacterial species in the sample. Slides were observed with an epifluorescence microscope (Axioplan 2; Zeiss, Sliedrecht, Netherlands), and images were acquired with a Zeiss MRM camera and compiled with the Zeiss microscopy image acquisition software (AxioVision version 4.7; Zeiss). The probes were subjected to excitation at 550 nm for Cy3, 494 for Fluos, and 649 nm for Cy5, and images were captured at emissions of 570 nm, 516 nm, and 670 nm, respectively.
A 250-ml volume of the culture broth was centrifuged for 10 min at 4,696 × g (Sorvall Legend X1R centrifuge; Thermo Fisher Scientific), the supernatant was discarded, and the pellet was washed with Tris-EDTA (TE; pH 8) buffer and stored at −20°C. DNA for library construction was extracted with the genomic DNA kit in combination with Genomic-tips 100/G (Qiagen Inc., CA, USA) according to the protocol, except for the addition of 2.6 mg ml−1 Zymolyase (20T; Amsbio, UK) and 4 mg ml−1 lysozyme. Cells were subsequently lysed by application of 2.9 × 105-kPa pressure with a French press (Constant Systems Ltd., UK). After purification with the genomic tip kit, the starting amount of DNA was quantified with the Qubit double-stranded DNA (dsDNA) HS assay kit (Thermo Fisher Scientific) and the quality of DNA was assessed with NanoDrop 2000. Sequencing was performed in-house on an Illumina MiSeq Sequencer (Illumina, San Diego, CA, USA), with MiSeq Reagent kit v3 with 2 × 300-bp read length, and the DNA libraries were prepared using the Nextera XT DNA sample preparation kit (Illumina). Quality trimming, adapter removal, and contaminant filtering of paired-end sequencing reads were performed using BBDUK (BBTOOLS version 37.17; BBMap [B. Bushnell, https://sourceforge.net/projects/bbmap/, unpublished]). Trimmed reads for all samples were coassembled using metaSPAdes v3.10.1 (74) at default settings. MetaSPAdes iteratively assembled the metagenome using kmer sizes 21, 33, 55, 77, 99, and 127. Reads were mapped back to the metagenome for each sample separately using the Burrows-Wheeler Aligner 0.7.15 (BWA), employing the “mem” algorithm (75). The generated sequence mapping files were handled and converted as needed using SAMtools 2.1 (76). Metagenome binning was performed using five different binning algorithms: BinSanity v0.2.5.9 (32), COCACOLA (33), CONCOCT (34), MaxBin 2.0 2.2.3 (35), and MetaBAT 2 2.10.2 (31). The five bin sets were supplied to DAS Tool 1.0 (77) for consensus binning to obtain the final optimized bins. The quality of the generated bins was assessed using CheckM 1.0.7 (36). The bins with the highest coverage were annotated by the RAST server for further analysis (37). For the identification of the dominant individual bins, the complete 16S rRNA genes were submitted to the SILVA database, release 132 (41).
We thank Marcel van den Broek for assisting with the bioinformatics, Dimitry Sorokin for assistance with the isolation, Ben Abbas for help with the 16S rRNA gene amplicon sequencing, Eveline van den Berg for critical reading of the manuscript, Maarten Verhoeven, Ioannis Papapetridis and Jasmine Bracher for fruitful discussions, and Orgaworld Nederland B.V. and the butcher in Est for supplying the inoculum.
This research was supported by the SIAM Gravitation Grant 024.002.002, the Netherlands Organization for Scientific Research.
L.C.V., M.C.M.V.L., A.J.A.V.M., and J.T.P. designed the experimental setup. L.C.V., M.C.M.V.L., and J.T.P. wrote the manuscript. M.V.T.H. assisted with testing the experimental setup of the bioreactors. L.C.V. did all the wet lab experiments, analyzed the 16S rRNA gene amplicon sequencing, extracted the DNA for the metagenomic analysis, and annotated and analyzed the metagenomic data. P.D.L.T.C. conducted the library preparation and sequencing of the enrichment cultures, and J.F. performed the bioinformatic analysis of the metagenomic data set. All authors read and approved the final manuscript.
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Department of Biotechnology, Delft University of Technology, Delft, Netherlands
Present address: Antonius J. A. van Maris, Department of Industrial Biotechnology, School of Engineering Sciences in Chemistry, Biotechnology and Health, KTH Royal Institute of Technology, AlbaNova University Centre, Stockholm, Sweden.
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