Galacturonate Metabolism in Anaerobic ChemostatEnrichment Cultures: Combined Fermentation andAcetogenesis by the Dominant sp. nov. “CandidatusGalacturonibacter soehngenii”

Laura C. Valk,a Jeroen Frank,b Pilar de la Torre-Cortés,a Max van ’t Hof,a Antonius J. A. van Maris,a* Jack T. Pronk,a

Mark C. M. van Loosdrechta

aDepartment of Biotechnology, Delft University of Technology, Delft, NetherlandsbSoehngen Institute of Anaerobic Microbiology, Radboud University Nijmegen, Nijmegen, Netherlands

ABSTRACT Agricultural residues such as sugar beet pulp and citrus peel are rich inpectin, which contains galacturonic acid as a main monomer. Pectin-rich residuesare underexploited as feedstocks for production of bulk chemicals or biofuels. Theanaerobic, fermentative conversion of D-galacturonate in anaerobic chemostat en-richment cultures provides valuable information toward valorization of these pectin-rich feedstocks. Replicate anaerobic chemostat enrichments, with D-galacturonate asthe sole limiting carbon source and inoculum from cow rumen content and rottingorange peels, yielded stable microbial communities, which were dominated by anovel Lachnospiraceae species, for which the name “Candidatus Galacturonibactersoehngenii” was proposed. Acetate was the dominant catabolic product, with for-mate and H2 as coproducts. The observed molar ratio of acetate and the combinedamounts of H2 and formate deviated significantly from 1, which suggested thatsome of the hydrogen and CO2 formed during D-galacturonate fermentation wasconverted into acetate via the Wood-Ljungdahl acetogenesis pathway. Indeed, met-agenomic analysis of the enrichment cultures indicated that the genome of “Candi-datus G. soehngenii” encoded enzymes of the adapted Entner-Doudoroff pathwayfor D-galacturonate metabolism as well as enzymes of the Wood-Ljungdahl pathway.The simultaneous operation of these pathways may provide a selective advantageunder D-galacturonate-limited conditions by enabling a higher specific ATP produc-tion rate and lower residual D-galacturonate concentration than would be possiblewith a strictly fermentative metabolism of this carbon and energy source.

IMPORTANCE This study on D-galacturonate metabolism by open, mixed-culture en-richments under anaerobic, D-galacturonate-limited chemostat conditions shows astable and efficient fermentation of D-galacturonate into acetate as the dominant or-ganic fermentation product. This fermentation stoichiometry and population analy-ses provide a valuable baseline for interpretation of the conversion of pectin-rich ag-ricultural feedstocks by mixed microbial cultures. Moreover, the results of this studyprovide a reference for studies on the microbial metabolism of D-galacturonate un-der different cultivation regimes.

KEYWORDS acetogenesis, enrichment culture, fermentation, galacturonate,metagenomics

Conversion of agricultural and food-processing residues by open, mixed microbialcommunities 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,

Received 6 June 2018 Accepted 27 June2018

Accepted manuscript posted online 29June 2018

Citation Valk LC, Frank J, de la Torre-Cortés P,van 't Hof M, van Maris AJA, Pronk JT, vanLoosdrecht MCM. 2018. Galacturonatemetabolism in anaerobic chemostatenrichment cultures: combined fermentationand acetogenesis by the dominant sp. nov.“Candidatus Galacturonibacter soehngenii.”Appl Environ Microbiol 84:e01370-18. https://doi.org/10.1128/AEM.01370-18.

Editor Haruyuki Atomi, Kyoto University

Copyright © 2018 American Society forMicrobiology. All Rights Reserved.

Address correspondence to Laura C. Valk,l.c.valk@tudelft.nl.

* Present address: Antonius J. A. van Maris,Department of Industrial Biotechnology,School of Engineering Sciences in Chemistry,Biotechnology and Health, KTH Royal Instituteof Technology, AlbaNova University Centre,Stockholm, Sweden.

MICROBIAL ECOLOGY

crossm

September 2018 Volume 84 Issue 18 e01370-18 aem.asm.org 1Applied and Environmental Microbiology

on April 5, 2020 by guest

http://aem.asm

.org/D

ownloaded from

fermentative metabolism of these residues can yield more-valuable products, includingorganic acids, solvents, and transport fuels. Production of such compounds in nona-septic open cultures has the potential to enable much lower capital and processingcosts than currently achievable with pure-culture processes (1–3).

Continuously operated, nonaseptic cultures enrich for microbial consortia that thriveunder the imposed conditions (1, 4). By carefully designing and optimizing reactorconfigurations and cultivation regimes, microbial populations that specifically andreproducibly yield desired product profiles can be enriched (5). Parameters such asculture pH, biomass retention time, and temperature, either in steady-state or dynamiccultivation regimes, are among the key parameters influencing population compositionand product profiles (6, 7). An additional decisive parameter in determining productformation by enrichment cultures is feedstock composition. Cell wall polymers andstorage carbohydrates represent the majority of the fermentable compounds in mostplant-derived waste streams. Hydrolysis of these polymers by extracellular microbialenzymes generates monomers, which are the actual substrates for anaerobic fermen-tation. Model studies with enrichment cultures on individual monomeric substrates cangenerate valuable information on the metabolic pathways and product profiles thatmay be encountered under industrially relevant conditions.

Previous studies on anaerobic fermentation of plant biomass monomers by contin-uous anaerobic enrichment cultures focused mainly on hexose and pentose sugarsreleased from the cellulose and hemicellulose fractions of plant biomass (8–10). How-ever, depending on the feedstock, other monomers can represent a significant or eventhe 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 thatcontains D-galacturonic acid as a main constituent (11–14). Large-volume citrus peelwaste streams originate from producing orange juice concentrates, which occursmainly in the United States and Brazil. Sugar beet pulp and apple pomace are producedpredominantly in Europe as waste streams of sugar beet and apple processing, respec-tively (15, 16). These pectin-rich agricultural residues are currently mostly dried and soldas 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-volumeresidues represent a potential and currently underexploited feedstock for production ofbulk chemicals or biofuels.

Intensively studied sugar substrates such as glucose, xylose, or arabinose can befermented to commodity products, including ethanol and lactate, at near-theoreticalyields. However, the currently documented biochemical pathways for galacturonatemetabolism do not enable high carbon and electron yields of these fermentationproducts (18–20). A major constraint in known pathways for dissimilation of galacturo-nate, which is more oxidized than glucose, involves cleavage of a C6 intermediate intotwo C3 moieties, eventually leading to the redox-neutral generation of two moles ofpyruvate from one mole of galacturonate (Fig. 1) (21–26). As a consequence, metabolicreactions beyond pyruvate need to be redox balanced, for example, by conversion ofpyruvate into equimolar amounts of acetyl coenzyme A (acetyl-CoA) and formate (Fig.1). Consistent with this observation, engineered microbial strains exhibit low carbonyields on galacturonate for compounds that are more reduced than pyruvate, such asethanol and lactate (11, 19, 20, 27).

Despite its ecological and industrial significance as a plant-derived carbonsource, fermentative conversion of galacturonate by anaerobic chemostat enrich-ment cultures has not been studied. The aim of this study was to characterizeproduct profiles and microbial populations of open, mixed-culture chemostat cul-tures fed with D-galacturonate as the sole limiting substrate and compare theseresults to cultures fed with the much better studied neutral sugar glucose (8).

Valk et al. Applied and Environmental Microbiology

September 2018 Volume 84 Issue 18 e01370-18 aem.asm.org 2

on April 5, 2020 by guest

http://aem.asm

.org/D

ownloaded from

RESULTSEnrichment of galacturonate-fermenting microbial cultures in anaerobic che-

mostats. To study the composition and physiology of anaerobic microbial enrichmentcultures on D-galacturonate, duplicate chemostat cultures were grown on a syntheticmedium in which this pectin monomer was the sole carbon and energy substrate. Bothbioreactors were started by adding anaerobic shake flask precultures on the samemedium, which had been inoculated with rotting orange peel and cow rumen content.The bioreactors were operated as batch cultures until all galacturonate was consumedand then were switched to continuous operation at a dilution rate of 0.13 h�1. Thisdilution rate and other parameters were chosen to enable direct comparison withprevious experiments in glucose-limited chemostat cultures (8). Subsequently, productconcentrations and carbon dioxide production were monitored over a period of 6weeks. Constant rates of CO2 production and of metabolite concentrations in thechemostats indicated that a “metabolic steady state” was reached after 2 weeks ofcontinuous cultivation (approximately 63 generations of the overall microbial popula-tion) (Fig. 2). In the steady-state cultures, the D-galacturonate concentration remainedbelow the detection limit of 0.1 mM, consistent with the medium composition, whichwas designed to make D-galacturonate the growth-limiting nutrient. After 1 month ofsteady-state operation (approximately 135 generations), five samples were taken overa period of 5 days for further physiological and metagenomic analysis.

Product profiles in chemostat enrichment cultures suggest the involvement ofacetogenesis in anaerobic galacturonate metabolism. To investigate galacturonate

FIG 1 Simplified scheme of pathways for fermentation of D-galacturonate to acetate and formate or toacetate, hydrogen, and carbon dioxide. The black lines indicate conversions that are possible, and graylines indicate conversions that are not possible when fermenting D-galacturonate based on redoxequivalents produced in the adapted Entner-Doudoroff pathway. The dashed lines represent multiple-step conversions, with the number of dashes independent of the number of conversions. Abbreviationsfor enzyme activities: Pfl, pyruvate-formate lyase (EC 2.3.1.54); PFOR, pyruvate:ferredoxin oxidoreductase(EC 1.2.7.1); Fhl, formate hydrogen lyase (EC 1.17.1.9 together with EC 1.12.1.2). Adapted from González-Cabaleiro et al. (26).

Anaerobic Galacturonate Enrichment Cultures Applied and Environmental Microbiology

September 2018 Volume 84 Issue 18 e01370-18 aem.asm.org 3

on April 5, 2020 by guest

http://aem.asm

.org/D

ownloaded from

metabolism, fermentation products were analyzed in the culture supernatants andoff-gas of the steady-state enrichment cultures. In both reactors, acetate was the maincatabolic product of D-galacturonate fermentation, with additional production of H2

and formate (Table 1). Average recoveries of carbon (92%) and electrons (95%) in thesteady-state cultures indicated that the major fermentation products were accountedfor (see Table S1 in the supplemental material).

In contrast to previously analyzed anaerobic chemostat enrichment experiments onglucose, which were grown under the same conditions (8) as used in the present study,

FIG 2 Concentrations of catabolic products during anaerobic enrichment experiments in galacturonate-limited chemostat cultures of bioreactors 1 (A) and 2 (B) with continuous cultivation starting at day (d)1. Acetate (black circles) and formate (open circles) concentrations are in millimolar; CO2 production(black squares) is in millimoles per hour per liter. Both bioreactors were operated at a dilution rate of 0.13h�1, with pH 8, and at 30°C.

TABLE 1 Comparison of product yields of anaerobic chemostat enrichment cultures ongalacturonate and glucosea

Product

Yieldb

Galacturonate

GlucosecBioreactor 1 Bioreactor 2

Biomass 0.16 � 0.01 0.16 � 0.02 0.21 � 0.01Succinate �0.01 0.02 � 0.01Acetate 0.53 � 0.02 0.53 � 0.02 0.16 � 0.02Butyrate �0.01 �0.01 0.30 � 0.04Ethanol �0.01 �0.01 0.08 � 0.01Formate 0.16 � 0.01 0.16 � 0.01 0.22 � 0.01H2 (mol Cmols�1) 0.01 � 0.00 0.02 � 0.00 0.05 � 0.01CO2 0.05 � 0.00 0.06 � 0.00H2 � formate (mol Cmols�1) 0.17 � 0.02 0.18 � 0.01 0.27 � 0.01Acetyl-CoA derivatives (mol Cmols�1)d 0.27 � 0.02 0.27 � 0.02 0.27 � 0.01aReplicate enrichment cultures (bioreactors 1 and 2) were grown on synthetic medium with galacturonate asthe limiting substrate, pH 8, at 30°C and at a dilution rate of 0.13 h�1.

bYields of biomass and products on galacturonate or glucose are expressed as Cmol Cmol�1 unless statedotherwise. Data represent averages � SD for five sequential measurements during steady state.

cData for glucose-limited cultures (grown under conditions identical to those of glucuronate-limitedenrichment) are from Temudo et al. (8).

dThe sum of the acetyl-CoA derivatives acetate, butyrate, and ethanol in moles of acetyl-CoA per Cmol ofsubstrate.

Valk et al. Applied and Environmental Microbiology

September 2018 Volume 84 Issue 18 e01370-18 aem.asm.org 4

on April 5, 2020 by guest

http://aem.asm

.org/D

ownloaded from

the galacturonate-grown enrichment cultures did not produce detectable amounts ofreduced fermentation products such as butyrate or ethanol (Table 1). Qualitatively, theformation of acetate, formate, CO2, and H2 as major fermentation products wasconsistent with previously described pathways for bacterial galacturonate metabolism(Fig. 1). However, in contrast to the product distribution expected from such pathwaysand to previous observations on glucose-grown chemostat enrichment cultures (8), thesum of the acetyl-CoA-derived products significantly differed from the combinedformate and H2 concentrations. In bioreactors 1 and 2, the combined yields of hydro-gen 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 observedmetabolite profiles would be consistent with a contribution of acetogenesis via theWood-Ljungdahl (WL) pathway, which enables conversion of fermentatively producedhydrogen (or alternatively, reduced ferredoxin derived from a pyruvate:ferredoxinoxidoreductase, EC 1.2.7.1) and carbon dioxide to acetate (equation 1). The combina-tion of acetogenesis with fermentative conversion of D-galacturonate into acetate via aclassical modified Entner-Doudoroff (ED) route (equation 2; Fig. 1) allows for conversionof D-galacturonate to acetate without net production of hydrogen (equation 3; Fig. 3).

2 H2 � 1 CO2 → 0.5 C2H4O2 � 1 H2O (1)

C6H10O7 � 1 H2O → 2 C2H4O2 � 2 H2 � 2 CO2 (2)

C6H10O7 → 2.5 C2H4O2 � 1 CO2 (3)

Based on the assumption that D-galacturonate catabolism in the enrichment cul-tures indeed reflected simultaneous D-galacturonate fermentation via an ED pathwayand acetogenesis via the WL pathway, the distribution of carbon and electrons overboth pathways can be easily estimated from measured rates of D-galacturonate con-

FIG 3 Metabolic model of D-galacturonate catabolism in anaerobic chemostat enrichment cultures. Metabolicfluxes were calculated by mass balancing based on a simplified Wood-Ljungdahl pathway converting four H2 andtwo CO2 into one acetate (42) and D-galacturonate metabolism via an adapted Entner-Doudoroff pathway forgalacturonate catabolism yielding two molecules of pyruvate (21–25). Redox equivalents have been representedas [H] and could both be derived from H2 as well as NADH. Biomass (x)-specific production rates, given in millimolesgramx

�1 hour�1, represent averages � standard deviations of five sequential measurements during steady statefrom both biological duplicate bioreactors.

Anaerobic Galacturonate Enrichment Cultures Applied and Environmental Microbiology

September 2018 Volume 84 Issue 18 e01370-18 aem.asm.org 5

on April 5, 2020 by guest

http://aem.asm

.org/D

ownloaded from

sumption and of hydrogen, formate, and acetate production in the steady-state cul-tures (three equations with two unknowns [28]). Based on these calculations (seecalculations in the supplemental material), the average specific rates of acetate forma-tion from pyruvate generated in the ED pathway (qacetate,ED) and of acetogenesis via theWL pathway (qacetate,WL) were estimated to be 6.9 � 0.4 mmol (g biomass)�1 h�1 and1.7 � 0.2 mmol (g biomass)�1 h�1, respectively. In such a scenario, 16% � 3% of theproduced hydrogen and formate generated from D-galacturonate would have beenused for acetogenesis (Fig. 3).

Population analysis of anaerobic D-galacturonate-fermenting enrichment cul-tures. To study microbial population dynamics during enrichment and to assess if thehypothesis outlined above was supported by the presence of acetogenic microorgan-isms, 16S-rRNA gene amplicon sequencing was performed on samples from bothbioreactors. The inoculum used to inoculate both bioreactors contained species relatedto the genera Clostridium sensu stricto and Lachnoclostridium as well as to unculturedLachnospiraceae (Fig. 4). After complete consumption of D-galacturonate in the batchphase preceding the continuous culture, bioreactor 1 was dominated by bacteriarelated to Raoultella and Yersinia, with the latter showing a 99% identity to Yersiniamassiliensis in the SILVA small subunit (SSU) database release 128. At the samesampling time, bioreactor 2 was dominated by bacteria related to Klebsiella andEnterobacter. Despite this difference in population composition, product profiles weresimilar for the two bioreactors at this time point, with acetate as the dominant organiccatabolic product and formate as the second major product (Fig. 2).

Although the microbial community compositions were different at the start of thecontinuous enrichment, similar populations developed during prolonged continuousoperation of the two bioreactors. In both reactors, a bacterium related to Lachnotaleabecame dominant, with a side population of a Klebsiella-related bacterium (Fig. 4). Thesteady-state communities in the two replicates were highly similar, indicating that thechosen cultivation conditions generated a selective advantage for these species andtheir conversions.

Community distribution determination by FISH analysis. To further quantify theabundance of dominant community members during the steady-state measurements,fluorescence in situ hybridization (FISH) analysis (29, 30) was performed (Fig. 5). Ageneral probe (EUB338) was used to stain all bacterial cells, while based on the resultsof 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 bacteriarelated to Lachnotalea and Klebsiella, respectively. Microorganisms belonging to theLachnospiraceae were found to be abundant in both reactors, with another abundantpopulation of bacteria belonging to the Enterobacteriaceae (Fig. 4 and 5). Theseobservations correlated with the identification of a Lachnotalea- and a Klebsiella-relatedbacterium, respectively, in the 16S rRNA gene analysis.

Metagenomic analysis of pathways and species in the enrichment cultures. Toinvestigate whether the enriched Lachnotalea-related bacterium harbored genes en-coding enzymes of the WL pathway and for DNA-based identification of the dominantmicroorganisms, a metagenomic analysis was performed on the steady-state cultures.DNA of the entire community was extracted from both steady-state enrichmentcultures 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 denovo assembly and metagenome binning-based sequence composition and differentialcoverage, 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%) inboth enrichments was identified to be a member of the Lachnospiraceae by analysis ofthe small subunit rRNA gene of the constructed genome. An identity of 93% was foundwithin the SILVA SSU database release 132 with the closest cultured relative, Lachno-

Valk et al. Applied and Environmental Microbiology

September 2018 Volume 84 Issue 18 e01370-18 aem.asm.org 6

on April 5, 2020 by guest

http://aem.asm

.org/D

ownloaded from

talea glycerini, indicating that the dominant microorganism belongs to a novel genuswithin the Lachnospiraceae family (40, 41). We propose the name “Candidatus Galac-turonibacter soehngenii” (Ga=.lac.tu.ro.ni.bac.ter. N.L. masc. n., galacturoni, referring tothe utilization of galacturonate; bacter, the equivalent of Gr. neut. n. bactrum, a rod orstaff, referring to the shape; soeh.ng.en=.ii., N.L. masc. n. gen., named after NicolaasSöhngen, first Ph.D. student in Delft University of Technology and pioneer in anaerobic

FIG 4 Population dynamics in anaerobic chemostat enrichment cultures on galacturonate. Genus level population compositions werederived from the 16S rRNA gene amplicon sequencing data, which make up �1% of the amplicon sequences. Population dynamicsin bioreactors 1 (A) and 2 (B). Both bioreactors were operated at a dilution rate of 0.13 h�1, with pH 8, and at 30°C.

Anaerobic Galacturonate Enrichment Cultures Applied and Environmental Microbiology

September 2018 Volume 84 Issue 18 e01370-18 aem.asm.org 7

on April 5, 2020 by guest

http://aem.asm

.org/D

ownloaded from

microbiology) for this novel microbe. The assembled and annotated genome of thisorganism harbored all structural genes for the enzymes of the canonic ED pathwaysfor D-galacturonate catabolism, as well as multiple signature genes encodingenzymes of the WL pathway for acetogenesis, including fhs, which encodesformate-tetrahydrofolate ligase (EC 6.3.4.3) 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 1.2.7.4), and acsBCD, which encodes the CO-methylating acetyl-CoA synthase complex (EC 2.1.3.169), were not identified.

A complete set of WL pathway genes were identified in the metagenomic data setwithin the genome of a species closely related to a Sporomusa species (99% identityfound in SILVA SSU database release 132), within a genus known for harboringhomoacetogenic bacteria (45). However, the genome of the Sporomusa species had acoverage of only 9-fold � 2-fold and a completeness of 80%. This coverage corre-sponds to approximately 1% � 0% of the total coverage. The other genome with a highcoverage (166-fold � 12-fold; completeness, 100%), was closely related to Klebsiellaoxytoca (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. TheK. oxytoca genome did not harbor any WL pathway genes but, in accordance withliterature (46), did contain all the genes for the adapted ED pathway for D-galacturonatemetabolism.

FIG 5 Fluorescence in situ hybridization (FISH) of anaerobic chemostat enrichment cultures. Culture samples taken at day 37 in twoparallel enrichment cultures were stained with a general probe for bacteria (EUB338, blue), a probe against members of the Lachno-spiraceae family (Lac435, red), and a probe against members of the Enterobacteriaceae family (ENT, green). Unstained (a, c) and stained(b, d) cells in bioreactors 1 (a, b) and 2 (c, d). Bars, 20 �m. Both bioreactors were operated at a dilution rate of 0.13 h�1, with pH 8, andat 30°C.

Valk et al. Applied and Environmental Microbiology

September 2018 Volume 84 Issue 18 e01370-18 aem.asm.org 8

on April 5, 2020 by guest

http://aem.asm

.org/D

ownloaded from

DISCUSSION

The identification of acetate as the dominant catabolic product of anaerobic che-mostat enrichment cultures on D-galacturonate (Table 1) is consistent with early studieson the metabolism of pectin-fermenting bacteria (47–49). The narrow product range ofthe currently described D-galacturonate-fermenting organisms is likely to be a conse-quence of the redox-neutral conversion of D-galacturonate to pyruvate via a modifiedEntner-Doudoroff (ED) pathway (Fig. 1) (18). In contrast, glucose fermentation, in whichoxidative glycolytic pathways generate reduced cofactors that are subsequently reoxi-dized by reduction of pyruvate and/or acetylphosphate, generates a wide diversity offermentation products. In enrichment cultures on glucose, performed under conditionsidentical to those used in the present study, a 1:2 molar ratio of butyrate to formate orhydrogen in such cultures was consistent with a classical Embden-Meyerhof glycolysisand 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 occurvia pyruvate formate-lyase (EC 1.2.7.1), phosphate acetyltransferase (EC 2.3.1.8), andacetate kinase (EC 2.7.2.1) (C6H10O7 ¡ 2 C2H4O2 � 2 HCOOH), with a potential furtherconversion of formate to hydrogen and CO2 by formate hydrogen-lyase (EC 1.12.1.2and EC 1.17.1.9) (Fig. 1) (26, 50, 51). However, since the 1:1 molar ratio of acetate toformate or hydrogen expected in such a scenario was not observed, we hypothesizedthat, in the chemostat enrichment cultures, acetogenesis via the Wood-Ljungdahl (WL)pathway converted some of the hydrogen and CO2 produced during conversion ofpyruvate to acetate.

The enrichment cultures contained different species suspected to be capable ofacetogenesis. Based on a sequence coverage estimate, a member of the Sporomusagenus, 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 responsiblefor the inferred rates of acetogenesis in the cultures, its specific activity would have toexceed previously reported rates of acetogenesis (52, 53) by at least 1 order ofmagnitude. This makes it likely that another bacterium was responsible for the majorityof the acetogenesis.

The dominance of “Candidatus G. soehngenii,” which was clearly shown in both the16S 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

TABLE 2 Genesa of the Wood-Ljungdahl and adapted Entner-Doudoroff pathways identified in the genome of “Candidatus G. soehngenii”

Pathway and gene product EC no. Gene E value

Wood-Ljungdahl pathwayHydrogen dehydrogenase 1.12.1.2 hoxF 6.00 � 10�33

Formate dehydrogenase 1.17.1.9 fdhA 6.00 � 10�33

Formate-tetrahydrofolate ligase 6.3.4.3 fhs 0.00Methenyl-tetrahydrofolate cyclohydrolase/methylene-tetrahydrofolate dehydrogenase 3.5.4.9 and 1.5.1.5 foldD 4.00 � 10�117

Methyl-tetrahydrofolate reductase 1.5.1.20 metF 0.005-Methyl-tetrahydrofolate:corrinoid/iron-sulfur protein methyltransferase 2.1.1.258 acsE 6.00 � 10�32

CO-methylating acetyl-CoA synthase 2.3.1.169 acsBCD —b

Carbon-monoxide dehydrogenase 1.2.7.4 cooS —

Adapted Entner-Doudoroff pathwayUronate isomerase 5.3.1.12 uxaC 0.00Tagaturonate reductase 1.1.1.58 uxaB 0.00Altronate dehydratase 4.2.1.7 uxaA 0.002-Dehydro-3-deoxygluconokinase 2.7.1.45 kdgK 2.00 � 10�135

2-Dehydro-3-deoxyphosphogluconate aldolase 4.1.2.14 kdgA 2.00 � 10�135

Pyruvate formate-lyase 2.3.1.54 pfl 2.00 � 10�91

Pyruvate:ferredoxin oxidoreductase 1.2.7.1 PFOR 0.00Phosphate acetyltransferase 2.3.1.8 pta 2.00 � 10�142

Acetate kinase 2.7.2.1 ackA 4.0 � 10�163

aGenes with the highest annotation found and their corresponding expected value (E value) are listed.b—, no homologues were identified.

Anaerobic Galacturonate Enrichment Cultures Applied and Environmental Microbiology

September 2018 Volume 84 Issue 18 e01370-18 aem.asm.org 9

on April 5, 2020 by guest

http://aem.asm

.org/D

ownloaded from

Gram-positive microorganisms (54, 55). We therefore assume that “Candidatus G.soehngenii,” while underrepresented in the FISH analysis, was indeed dominant andresponsible for the main conversions observed in both bioreactors.

After the onset of continuous cultivation, “Candidatus G. soehngenii” graduallyreplaced other microorganisms (Fig. 3). The genome of this organism, which becamedominant at the end of duplicate chemostat enrichment experiments, harbored mostgenes of the WL pathway. However, we did not identify genes with a high sequencesimilarity to known cooS and acsBCD genes respectively encoding carbon-monoxidedehydrogenase (EC 1.2.7.4) and CO-methylating acetyl-CoA synthase complex subunits(EC 2.1.3.169) (42). Genomes of Lachnospiraceae species remain underexplored andcontain many nonannotated genes, and the possibility of finding novel functionalhomologues of the Wood-Ljungdahl pathway might be interesting for future research(56). Alternatively, the “missing” cooS and acsBCD genes may be an artifact of themetagenome assembly and analysis. All traditional methods were used in an attemptto isolate “Candidatus G. soehngenii” (for a detailed list of isolation methods, see TableS2 in the supplemental material), but none was successful. We postulate that “Candi-datus G. soehngenii” is very well adapted to growth at limiting substrate concentrations(K-strategist) as occurring in a chemostat-operated bioreactor (57). Alternative isolationprocedures 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 ace-togenesis.

Microbial competition is often interpreted in terms of the affinity for a singlegrowth-limiting nutrient (qs

max/Ks) (58). Under nutrient-limited conditions, simultane-ous utilization of mixed substrates enables growth at lower concentrations of each ofthe 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 galacturonateconcentrations in the chemostat cultures, thereby conferring a selective advantage to“Candidatus G. soehngenii.” Additionally, the net ATP yield of D-galacturonate fermen-tation via an ED-type pathway amounts to 3 mol ATP (mol galacturonate)�1 or 1.5 molATP (mol acetate)�1, while the supplemental formation of acetate via the WL pathwayis estimated to yield an additional 0.33 mol ATP (mol acetate)�1, assuming chemios-motic energy conservation due to the generation of a transmembrane electrochemicalion gradient (61). With a constant growth rate under chemostat operations, the totalATP flux, qATP, required for growth will also remain constant. When the ATP is suppliedby both the adapted ED pathway and the WL pathway, the relative qATP derived fromthe ED pathway can decrease, without lowering the overall qATP. Lowering the fluxthrough the ED pathway decreases the galacturonate biomass specific uptake rate (qs),and in accordance to the substrate uptake kinetics, shown in equation 4, this willdecrease the galacturonate concentration (Cs) in the bioreactor (62).

qs � qs,max ·Cs

Cs � Ks(4)

The very slow replacement of nonacetogenic bacteria in the chemostat enrichmentcultures is in line with the relatively small impact of acetogenesis on the kinetics andenergetics of D-galacturonate metabolism. Moreover, the dilution rate of the chemostatcultures was at the upper end of specific growth rates reported for typical hetero-trophic homoacetogens on (semi-) defined media (63–65). Additionally, sparging of thereactors with nitrogen gas may have stripped H2 formed by D-galacturonate fermen-tation. 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 WLpathway (for calculations, see the supplemental material) (66, 67). With the Gibbs freeenergy change close to the minimal driving force for catabolic reactions, this low in situhydrogen partial pressure may have limited the rate of H2 oxidation in acetogenesis.This interpretation is consistent with the dominance of fast-growing nonacetogenic

Valk et al. Applied and Environmental Microbiology

September 2018 Volume 84 Issue 18 e01370-18 aem.asm.org 10

on April 5, 2020 by guest

http://aem.asm

.org/D

ownloaded from

bacteria (e.g., Klebsiella and Clostridium species) during the batch phase that precededthe chemostat enrichment.

Although involvement of the WL pathway has been demonstrated in anaerobicchemostat 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 operationalconditions similar to those used in this study [8]). This observation suggests thatpotentially lowering the residual glucose concentration by simultaneous acetogenesisdid not provide a selective advantage in anaerobic enrichment cultures fed withglucose. More research is required to elucidate this difference in usage of the Wood-Ljungdahl pathway in glucose- and galacturonate-limited cultures. Although “Candida-tus G. soehngenii,” which became dominant in the D-galacturonate-limited enrichmentcultures in this study, differed from the Clostridium quinii that dominated similarcultures grown on glucose as the carbon source (69), both are members of theClostridiales. This observation illustrates the importance of these organisms in fermen-tative conversion of carbohydrates and related compounds in carbon-limited, anaero-bic environments (69).

MATERIALS AND METHODSReactor operation. 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 anaerobicconditions. The pH was controlled at pH 8 � 0.1 by automatic titration (ADI 1030 Bio controller) with a1 M NaOH solution. The dilution rate was 0.13 h�1, and the working volume was kept constant byperistaltic effluent pumps (Masterflex; Cole-Parmer, Vernon Hills, IL, USA) coupled to electrical levelsensors.

Steady-state characterization. To characterize the product spectrum, the reactor was run incontinuous mode until stable product composition and biomass concentration were established. Steadystate was reached after 2 weeks (63 generations). On-line analysis of system stability was achieved byonline monitoring of the gas productivities, in the form of CO2, and base addition. On-line gas detectionof CO2 was done with a Rosemount Analytical NGA 2000 MLT 1 multicomponent analyzer (infrareddetector). 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 trendshowing increase or decrease, a set of samples were taken during the subsequent cycles. The concen-trations of soluble organic fermentation products, hydrogen produced, and biomass in the reactorvolume were determined.

Inoculum. The reactor was inoculated (1%, vol/vol) with open mixed precultures started with amixture of two different sources. The first inoculum was obtained from the content of rumen offal ofa grass-fed cow (Est, Netherlands). The second inoculum was a sample from organic waste containing alarge amount of citrus peels (Orgaworld Nederland B.V., Netherlands). Shake flasks were inoculated induplicate with compost (2%, vol/vol) and rumen extract (2% vol/vol), and the initial pH was adjusted to8 with a 2 M KOH solution. The shake flasks were cultivated in an anaerobic chamber (Bactron III; ShelLab, Cornelius, USA) at 30°C with a gas composition of 89% N2, 6% CO2, and 5% H2.

Medium. 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 wereprepared and fed separately. The D-galacturonate solution was sterilized by filtration (0.2 �m MediakapPlus; Spectrum Laboratories, Rancho Dominguez, CA, USA), and the mineral solution was autoclaved for20 min at 121°C. Three milliliters PluronicPE 6100 antifoam (BASF, Ludwigshafen, Germany) was addedper 40 liters mineral solution to avoid excessive foaming.

Analytical methods of substrate and extracellular metabolites. To determine substrate andextracellular metabolite concentrations, reactor sample supernatant was obtained by centrifugation ofculture samples (Biofuge Pico; Heraeus, Hanau, Germany) and subsequent filtration (0.2 �m Millex-HV;Millipore-Merck, Darmstadt, Germany). Concentrations of galacturonate and extracellular metaboliteswere 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 mlmin�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; ThermoScientific, USA).

Biomass. Culture dry weight was measured by filtering 20 ml of culture broth over predried andpreweighed membrane filters (0.2-�m Supor-200; Pall corporation, New York, NY, USA), which were thenwashed with demineralized water, dried in a microwave oven (Easytronic M591; Whirlpool, BentonHarbor, MI, USA), and weighed again. Carbon balances and electron balances were established based onthe number of carbon atoms and electrons per mole, while a standard biomass composition ofCH1.8O0.5N0.2 was assumed (70).

Anaerobic Galacturonate Enrichment Cultures Applied and Environmental Microbiology

September 2018 Volume 84 Issue 18 e01370-18 aem.asm.org 11

on April 5, 2020 by guest

http://aem.asm

.org/D

ownloaded from

Microbial community stability analysis. The bacterial DNA for the community analysis over timewas obtained by extracting 2 ml broth and pelleting the biomass before storing it at �80°C for furtheranalysis. The DNA was extracted using the UltraClean DNA isolation kit (MOBIO laboratories Inc., USA) asper the manufacturer’s instructions. The 16S rRNA genes of days 1 and 38 of bioreactor 1 and day 38 ofbioreactor 2 were analyzed using amplicon sequencing with an Illumina HiSeq (Novogene BioinformaticsTechnology 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 accord-ing to the method of van den Berg et al. (71). Representative sequences for each operational taxonomicunit were submitted for BLAST analysis under default conditions. The extracted genomic DNAs of theinoculum, from days 9 and 14 of bioreactor 1 and days 1, 9, and 14 of bioreactor 2, were subsequentlyused for a two-step PCR targeting the 16S rRNA gene of most bacteria and archaea. For this, the primersU515F (5=-GTGYCAGCMGCCGCGGTA-3=) and U1071R (5=-GARCTGRCGRCRRCCATGCA-3=) were used ac-cording to the method of Wang et al. (72). The first amplification was performed to enrich for 16S rRNAgenes, using a quantitative PCR assay with 2� iQ SYBR green Supermix (Bio-Rad, CA, USA), 500 nMprimers each, and 1 to 50 ng of genomic DNA was added per well to a final volume of 20 �l. Thethermocycles consisted of a first denaturation at 95°C for 5 min, 20 cycles at 95°C for 30 s, 50°C for 40s, and 72°C for 40 s, and a final extension of 72°C for 7 min. 454-adapters (Roche) and MID tags at theU515F primer were added to the produced PCR products. The second amplification was similar to thefirst one, with the exception of the use of Taq PCR master mix (Qiagen Inc., CA, USA); the program wasrun for 15 cycles, and the template (the product from step one) was diluted 10 times. After the secondamplification, the PCR products were pooled equimolar and purified over an agarose gel using a GeneJETgel extraction kit (Thermo Fisher Scientific, Netherlands). The resulting library was sent for 454 sequenc-ing and run in 1/8 lane with titanium chemistry by Macrogen Inc. (Seoul, South Korea). After analysis, thereads 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 123.1SSURef 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 andspecies abundance.

Fluorescence in situ hybridization. FISH was performed as described in reference 73, using ahybridization buffer containing 25% (vol/vol) formamide. Probes were synthesized and 5= labeled eitherwith 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 eubacterialspecies 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 theZeiss microscopy image acquisition software (AxioVision version 4.7; Zeiss). The probes were subjectedto excitation at 550 nm for Cy3, 494 for Fluos, and 649 nm for Cy5, and images were captured atemissions of 570 nm, 516 nm, and 670 nm, respectively.

Metagenomics. 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 pelletwas washed with Tris-EDTA (TE; pH 8) buffer and stored at �20°C. DNA for library construction wasextracted 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 mgml�1 lysozyme. Cells were subsequently lysed by application of 2.9 � 105-kPa pressure with a Frenchpress (Constant Systems Ltd., UK). After purification with the genomic tip kit, the starting amount of DNAwas quantified with the Qubit double-stranded DNA (dsDNA) HS assay kit (Thermo Fisher Scientific) andthe quality of DNA was assessed with NanoDrop 2000. Sequencing was performed in-house on anIllumina MiSeq Sequencer (Illumina, San Diego, CA, USA), with MiSeq Reagent kit v3 with 2 � 300-bpread 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 readswere performed using BBDUK (BBTOOLS version 37.17; BBMap [B. Bushnell, https://sourceforge.net/projects/bbmap/, unpublished]). Trimmed reads for all samples were coassembled using metaSPAdesv3.10.1 (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 usingthe Burrows-Wheeler Aligner 0.7.15 (BWA), employing the “mem” algorithm (75). The generated se-quence mapping files were handled and converted as needed using SAMtools 2.1 (76). Metagenomebinning 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 DASTool 1.0 (77) for consensus binning to obtain the final optimized bins. The quality of the generated binswas assessed using CheckM 1.0.7 (36). The bins with the highest coverage were annotated by the RASTserver for further analysis (37). For the identification of the dominant individual bins, the complete 16SrRNA genes were submitted to the SILVA database, release 132 (41).

TABLE 3 FISH probes used in this study

Probe Dye Sequence (5=–3=) Specificity Reference(s)

ENT Fluos CCCCCWCTTTGGTCTTGCa Enterobacteriaceae exceptProteus spp.

78

Lac435 CY3 TCTTCCCTGCTGATAGA Lachnospiraceae family 79EUB338 CY5 GCTGCCTCCCGTAGGAGT Most bacteria 80, 81aW stands for A or T.

Valk et al. Applied and Environmental Microbiology

September 2018 Volume 84 Issue 18 e01370-18 aem.asm.org 12

on April 5, 2020 by guest

http://aem.asm

.org/D

ownloaded from

Availability of data. Genome sequence data of the amplicon sequences of the 16S rRNA geneanalysis have been deposited at the NCBI GenBank (https://www.ncbi.nlm.nih.gov/GenBank/) underaccession numbers MG982381 to MG982451. Genome sequence data of the metagenomic assembly ofthe galacturonate-fermenting enrichment culture metagenome, biosample SAMN08337232, have beendeposited in the NCBI genomic database (www.ncbi.nlm.nih.gov/genome/) under BioProject IDPRJNA429181.

SUPPLEMENTAL MATERIAL

Supplemental material for this article may be found at https://doi.org/10.1128/AEM.01370-18.

SUPPLEMENTAL FILE 1, PDF file, 0.6 MB.

ACKNOWLEDGMENTSWe 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 ampliconsequencing, Eveline van den Berg for critical reading of the manuscript, MaartenVerhoeven, Ioannis Papapetridis and Jasmine Bracher for fruitful discussions, andOrgaworld 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 Neth-erlands 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 experi-mental setup of the bioreactors. L.C.V. did all the wet lab experiments, analyzed the 16SrRNA gene amplicon sequencing, extracted the DNA for the metagenomic analysis, andannotated and analyzed the metagenomic data. P.D.L.T.C. conducted the library prep-aration and sequencing of the enrichment cultures, and J.F. performed the bioinfor-matic analysis of the metagenomic data set. All authors read and approved the finalmanuscript.

REFERENCES1. Kleerebezem R, van Loosdrecht MCM. 2007. Mixed culture biotechnol-

ogy for bioenergy production. Curr Opin Biotechnol 18:207–212. https://doi.org/10.1016/j.copbio.2007.05.001.

2. Angenent LT, Karim K, Al-Dahhan MH, Wrenn BA, Domiguez-Espinosa R.2004. Production of bioenergy and biochemicals from industrial andagricultural wastewater. Trends Biotechnol 22:477– 485. https://doi.org/10.1016/j.tibtech.2004.07.001.

3. Claassen PAM, van Lier JB, Lopez Contreras AM, van Niel EWJ, Sijtsma L,Stams AJM, de Vries SS, Weusthuis RA. 1999. Utilisation of biomass forthe supply of energy carriers. Appl Microbiol Biotechnol 52:741–755.https://doi.org/10.1007/s002530051586.

4. Beijerinck MW. 1901. Enrichment culture studies with urea bacteria.Zentralbl Bakteriol 1901(Part II):33– 61.

5. Rodríguez J, Kleerebezem R, Lema JM, van Loosdrecht MCM. 2006.Modeling product formation in anaerobic mixed culture fermentations.Biotechnol Bioeng 93:592– 606. https://doi.org/10.1002/bit.20765.

6. Zoetemeyer RJ, Matthijsen AJCM, Cohen A, Boelhouwer C. 1982. Productinhibition in the acid forming stage of the anaerobic digestion process.Water Res 16:633– 639. https://doi.org/10.1016/0043-1354(82)90084-7.

7. Lengeler JW, Drews G, Schlegel HG. 1998. Biology of prokaryotes. Black-well, Oxford, England.

8. Temudo MF, Mato T, Kleerebezem R, van Loosdrecht MCM. 2009. Xyloseanaerobic conversion by open-mixed cultures. Appl Microbiol Biotech-nol 82:231–239. https://doi.org/10.1007/s00253-008-1749-y.

9. Lin CY, Cheng CH. 2006. Fermentative hydrogen production from xyloseusing anaerobic mixed microflora. Int J Hydrogen Energy 31:832– 840.https://doi.org/10.1016/j.ijhydene.2005.08.010.

10. Zhang F, Yang JH, Dai K, Chen Y, Li QR, Gao FM, Zeng RJ. 2016.Characterization of microbial compositions in a thermophilic chemostatof mixed culture fermentation. Appl Microbiol Biotechnol 100:1511–1521. https://doi.org/10.1007/s00253-015-7130-z.

11. Grohmann K, Manthey JA, Cameron RG, Buslig BS. 1998. Fermentation ofgalacturonic acid and pectin-rich materials to ethanol by geneticallymodified strains of Erwinia. Biotechnol Lett 20:195–200. https://doi.org/10.1023/A:1005349112770.

12. Richard P, Hilditch S. 2009. D-Galacturonic acid catabolism in microor-ganisms and its biotechnological relevance. Appl Microbiol Biotechnol82:597– 604. https://doi.org/10.1007/s00253-009-1870-6.

13. Mohnen D. 2008. Pectin structure and biosynthesis. Curr Opin Plant Biol11:266 –277. https://doi.org/10.1016/j.pbi.2008.03.006.

14. Grohmann K, Bothast RJ. 1994. Pectin-rich residues generated by pro-cessing of citrus fruits, apples, and sugar beets— enzymatic hydrolysisand biological conversion to value-added products. ACS Symp Ser En-zym Convers Biomass Fuels Prod 566:372–390. https://doi.org/10.1021/bk-1994-0566.ch019.

15. Bayer AG. Why sugar beet matters to us. https://www.cropscience.bayer.com/en/crop-compendium/key-crops/sugar-beets. Accessed 15 January2018.

16. US Department of Agriculture. 2017. Citrus: world markets and trade.https://apps.fas.usda.gov/psdonline/circulars/citrus.pdf.

17. Food and Agriculture Organization of the United Nations. 2016. Crops,production quantities of sugar beet, citrus fruits and apples in the world.FAOSTAT. http://www.fao.org/faostat/en/#data/QC.

18. van Maris AJA, Abbott DA, Bellissimi E, van den Brink J, Kuyper M, LuttikMAH, Wisselink HW, Scheffers WA, van Dijken JP, Pronk JT. 2006. Alco-holic fermentation of carbon sources in biomass hydrolysates by Sac-charomyces cerevisiae: current status. Antonie Van Leeuwenhoek 90:391– 418. https://doi.org/10.1007/s10482-006-9085-7.

19. Doran JB, Cripe J, Sutton M, Foster B. 2000. Fermentations of pectin-richbiomass with recombinant bacteria to produce fuel ethanol. ApplBiochem Biotechnol 84 – 86:141–152.

20. Doran-Peterson J, Cook DM, Brandon SK. 2008. Microbial conversion ofsugars from plant biomass to lactic acid or ethanol. Plant J 54:582–592.https://doi.org/10.1111/j.1365-313X.2008.03480.x.

21. Ashwell G, Wahba AJ, Hickman J. 1960. Uronic acid metabolism inbacteria I purification and properties of uronic acid isomerase in Esche-richia coli. J Biol Chem 235:1559 –1565.

22. Hickman J, Ashwell G. 1960. Uronic acid metabolism in bacteria. II.Purification and properties of D-altronic acid and D-mannonic aciddehydrogenases in Escherichia coli. J Biol Chem 235:1566 –1570.

Anaerobic Galacturonate Enrichment Cultures Applied and Environmental Microbiology

September 2018 Volume 84 Issue 18 e01370-18 aem.asm.org 13

on April 5, 2020 by guest

http://aem.asm

.org/D

ownloaded from

23. Smiley JD, Ashwell G. 1960. Uronic acid metabolism in bacteria. III.Purification and properties of D-altronic acid and D-mannonic aciddehydrases in Escherichia coli. J Biol Chem 235:1571–1575.

24. Cynkin MA, Ashwell G. 1960. Uronic acid metabolism in bacteria. IV.Purification and properties of 2-keto-3-deoxy-D-gluconokinase in Esch-erichia coli. J Biol Chem 235:1576 –1579.

25. Kovachevich R, Wood WA. 1955. Carbohydrate metabolism by Pseu-domonas fluorescens IV purification and properties of 2-keto-3-deoxy-6-phospho-gluconate aldolase. J Biol Chem 213:757–767.

26. González-Cabaleiro R, Lema JM, Rodríguez J. 2015. Metabolic energy-based modelling explains product yielding in anaerobic mixed cul-ture fermentations. PLoS One 10:e0126739. https://doi.org/10.1371/journal.pone.0126739.

27. Grohmann K, Baldwin EA, Buslig BS. 1994. Fermentation of galacturonicacid and other sugars in orange peel hydrolysates by the ethanologenicstrain of Escherichia coli. Biotechnol Lett 16:281–286. https://doi.org/10.1007/BF00134626.

28. Noorman HJ, Heijnen JJ, Luyben KCAM. 1991. Linear relations in micro-bial reaction systems: a general overview of their origin, form, and use.Biotechnol Bioeng 38:603– 618. https://doi.org/10.1002/bit.260380606.

29. Brooks JP, Edwards DJ, Harwich MD, Jr, Rivera MC, Fettweis JM, SerranoMG, Reris RA, Sheth NU, Huang B, Girerd P, Vaginal Microbiome Con-sortium, Strauss JF, III, Jefferson KK, Buck GA. 2015. The truth aboutmetagenomics: quantifying and counteracting bias in 16S rRNA studies.BMC Microbiol 15:66. https://doi.org/10.1186/s12866-015-0351-6.

30. Kembel SW, Wu M, Eisen JA, Green JL. 2012. Incorporating 16S genecopy number information improves estimates of microbial diversity andabundance. PLoS Comput Biol 8:16 –18. https://doi.org/10.1371/journal.pcbi.1002743.

31. Kang DD, Froula J, Egan R, Wang Z. 2015. MetaBAT, an efficient tool foraccurately reconstructing single genomes from complex microbial com-munities. PeerJ 3:e1165. https://doi.org/10.7717/peerj.1165.

32. Graham ED, Heidelberg JF, Tully BJ. 2017. BinSanity: unsupervised clus-tering of environmental microbial assemblies using coverage and affin-ity propagation. PeerJ 5:e3035. https://doi.org/10.7717/peerj.3035.

33. Lu YY, Chen T, Fuhrman JA, Sun F, Sahinalp C. 2017. COCACOLA: binningmetagenomic contigs using sequence COmposition, read CoverAge,CO-alignment and paired-end read LinkAge. Bioinformatics 33:791–798.https://doi.org/10.1093/bioinformatics/btw290.

34. Alneberg J, Bjarnason BS, de Bruijn I, Schirmer M, Quick J, Ijaz UZ, LahtiL, Loman NJ, Andersson AF, Quince C. 2014. Binning metagenomiccontigs by coverage and composition. Nat Methods 11:1144 –1146.https://doi.org/10.1038/nmeth.3103.

35. Wu Y-W, Simmons BA, Singer SW. 2016. MaxBin 2.0: an automated binningalgorithm to recover genomes from multiple metagenomic datasets. Bioin-formatics 32:605–607. https://doi.org/10.1093/bioinformatics/btv638.

36. Parks DH, Imelfort M, Skennerton CT, Hugenholtz P, Tyson GW. 2015.CheckM: assessing the quality of microbial genomes recovered fromisolates, single cells, and metagenomes. Genome Res 25:1043–1055.https://doi.org/10.1101/gr.186072.114.

37. Aziz RK, Bartels D, Best AA, DeJongh M, Disz T, Edwards RA, Formsma K,Gerdes S, Glass EM, Kubal M, Meyer F, Olsen GJ, Olson R, Osterman AL,Overbeek RA, McNeil LK, Paarmann D, Paczian T, Parrello B, Pusch GD,Reich C, Stevens R, Vassieva O, Vonstein V, Wilke A, Zagnitko O. 2008.The RAST server: rapid annotations using subsystems technology. BMCGenomics 9:75. https://doi.org/10.1186/1471-2164-9-75.

38. Overbeek R, Olson R, Pusch GD, Olsen GJ, Davis JJ, Disz T, Edwards RA,Gerdes S, Parrello B, Shukla M, Vonstein V, Wattam AR, Xia F, Stevens R.2014. The SEED and the Rapid Annotation of microbial genomes usingSubsystems Technology (RAST). Nucleic Acids Res 42:206 –214. https://doi.org/10.1093/nar/gkt1226.

39. Brettin T, Davis JJ, Disz T, Edwards RA, Gerdes S, Olsen GJ, Olson R,Overbeek R, Parrello B, Pusch GD, Shukla M, Thomason JA, Stevens R,Vonstein V, Wattam AR, Xia F. 2015. RASTtk: a modular and extensibleimplementation of the RAST algorithm for building custom annotationpipelines and annotating batches of genomes. Sci Rep 5:8365. https://doi.org/10.1038/srep08365.

40. Yarza P, Yilmaz P, Pruesse E, Glöckner FO, Ludwig W, Schleifer KH,Whitman WB, Euzéby J, Amann R, Rosselló-Móra R. 2014. Uniting theclassification of cultured and uncultured bacteria and archaea using 16SrRNA gene sequences. Nat Rev Microbiol 12:635– 645. https://doi.org/10.1038/nrmicro3330.

41. Pruesse E, Peplies J, Glöckner FO. 2012. SINA: accurate high-throughput

multiple sequence alignment of ribosomal RNA genes. Bioinformatics28:1823–1829. https://doi.org/10.1093/bioinformatics/bts252.

42. Ragsdale SW. 2008. Enzymology of the Wood-Ljungdahl pathway ofacetogenesis. Ann N Y Acad Sci 1125:129 –136. https://doi.org/10.1196/annals.1419.015.

43. Henderson G, Naylor GE, Leahy SC, Janssen PH. 2010. Presence of novel,potentially homoacetogenic bacteria in the rumen as determined byanalysis of formyltetrahydrofolate synthetase sequences from rumi-nants. Appl Environ Microbiol 76:2058 –2066. https://doi.org/10.1128/AEM.02580-09.

44. Gagen EJ, Denman SE, Padmanabha J, Zadbuke S, Jassim Al R, MorrisonM, McSweeney CS. 2010. Functional gene analysis suggests differentacetogen populations in the bovine rumen and tammar wallaby fores-tomach. Appl Environ Microbiol 76:7785–7795. https://doi.org/10.1128/AEM.01679-10.

45. Breznak JA. 2006. The genus Sporomusa, p 991–1001. In Dworkin M,Falkow S, Rosenberg E, Schleifer KH, Stackebrandt E (ed), The pro-karyotes, vol 4. Bacteria: Firmicutes, Cyanobacteria, 3rd ed. Springer-Verlag, New York, NY.

46. Shin SH, Kim S, Kim JY, Lee S, Um Y, Oh MK, Kim YR, Lee J, Yang KS. 2012.Complete genome sequence of Klebsiella oxytoca KCTC 1686, used inproduction of 2,3-butanediol. J Bacteriol 194:2371–2372. https://doi.org/10.1128/JB.00026-12.

47. Ng H, Vaughn RH. 1963. Clostridium rubrum sp. n. and other pectinolyticClostridia from soil. J Bacteriol 85:1104 –1113.

48. Burkey LA. 1928. The fermentation of cornstalks and their constituents.I. Studies on the pectin-fermenting bacteria. Retrospective theses anddissertations, 14219. Iowa State College, Ames, IA.

49. Lanigan GW. 1959. Studies on the pectinolytic anaerobes Clostridiumflavum and Clostridium laniganii. J Bacteriol 77:1–9.

50. Thauer RK, Jungermann K, Decker K. 1977. Energy conservation inchemotrophic anaerobic bacteria. Bacteriol Rev 41:100 –180.

51. Thauer RK, Kirchniawy FH, Jungermann KA. 1972. Properties and func-tion of the pyruvate formate-lyase reaction in Clostridiae. Eur J Biochem27:282–290. https://doi.org/10.1111/j.1432-1033.1972.tb01837.x.

52. Valentine DL, Sessions AL, Tyler SC, Chidthaisong A. 2004. Hydrogenisotope fractionation during H2/CO2 acetogenesis: hydrogen utilizationefficiency and the origin of lipid-bound hydrogen. Geobiology2:179 –188. https://doi.org/10.1111/j.1472-4677.2004.00030.x.

53. Tremblay P, Höglund D, Koza A, Bonde I, Zhang T. 2015. Adaptation ofthe autotrophic acetogen Sporomusa ovata to methanol accelerates theconversion of CO2 to organic products. Sci Rep 5:161168. https://doi.org/10.1038/srep16168.

54. Franks AH, Harmsen HJM, Gerwin C, Jansen GJ, Schut F, Gjalt W. 1998.Variations of bacterial populations in human feces measured by fluo-rescent in situ hybridization with group-specific 16S rRNA-targeted oli-gonucleotide probes. Appl Environ Microbiol 64:3336 –3345.

55. Moter A, Göbel UB. 2000. Fluorescence in situ hybridization (FISH) fordirect visualization of microorganisms. J Microbiol Methods 41:85–112.https://doi.org/10.1016/S0167-7012(00)00152-4.

56. Meehan CJ, Beiko RG. 2014. A phylogenomic view of ecological specializa-tion in the Lachnospiraceae, a family of digestive tract-associated bacteria.Genome Biol Evol 6:703–713. https://doi.org/10.1093/gbe/evu050.

57. Andrews JH, Harris RF. 1986. r- and K-selection and microbial ecology.Adv Microb Ecol 9:99 –147. https://doi.org/10.1007/978-1-4757-0611-6_3.

58. Kuenen JG. 2015. Continuous cultures (chemostats). Ref Module BiomedRes 1–19. https://doi.org/10.1016/B978-0-12-801238-3.02490-9.

59. Lendenmann U, Snozzi M, Egli T. 1996. Kinetics of the simultaneousutilization of sugar mixtures by Escherichia coli in continuous culture.Appl Environ Microbiol 62:1493–1499.

60. Egli T, Lendenmann U, Snozzi M. 1993. Kinetics of microbial growth withmixtures of carbon sources. Antonie Van Leeuwenhoek 63:289 –298.https://doi.org/10.1007/BF00871224.

61. Schuchmann K, Müller V. 2014. Autotrophy at the thermodynamic limitof life: a model for energy conservation in acetogenic bacteria. Nat RevMicrobiol 12:809 – 821. https://doi.org/10.1038/nrmicro3365.

62. Monod J. 1942. Recherches sur la croissance des cultures bacteriennes.Hermann et Cie, Paris, France.

63. Savage MD, Drake HL. 1986. Adaptation of the acetogen Clostridium ther-moautotrophicum to minimal medium. J Bacteriol 165:315–318. https://doi.org/10.1128/jb.165.1.315-318.1986.

64. Daniel SL, Hsu T, Dean SI, Drake HL, Ljungdahl L. 1990. Characterizationof the H2- and CO-dependent chemolithotrophic potentials of the ace-

Valk et al. Applied and Environmental Microbiology

September 2018 Volume 84 Issue 18 e01370-18 aem.asm.org 14

on April 5, 2020 by guest

http://aem.asm

.org/D

ownloaded from

togens Clostridium thermoaceticum and Acetogenium kivuit. J Bacteriol172:4464 – 4471. https://doi.org/10.1128/jb.172.8.4464-4471.1990.

65. Lundie LL, Jr, Drake HL. 1984. Development of a minimally definedmedium for the acetogen Clostridium thermoaceticum. J Bacteriol 159:700 –703.

66. De Kok S, Meijer J, van Loosdrecht MCM, Kleerebezem R. 2013. Impactof dissolved hydrogen partial pressure on mixed culture fermenta-tions. Appl Microbiol Biotechnol 97:2617–2625. https://doi.org/10.1007/s00253-012-4400-x.

67. Heijnen JJ. 1999. Bioenergetics of microbial growth, p 267–291. InFlickinger MC, Drew SW (ed), Encyclopedia of bioprocess technology:fermentation. biocatalysis, and bioseparation. John Wiley & Sons, Hobo-ken, NJ.

68. Ragsdale SW, Pierce E. 2008. Acetogenesis and the Wood-Ljungdahlpathway of CO2 fixation. Biochim Biophys Acta 1784:1873–1898. https://doi.org/10.1016/j.bbapap.2008.08.012.

69. Temudo MF, Muyzer G, Kleerebezem R, van Loosdrecht MCM. 2008.Diversity of microbial communities in open mixed culture fermentations:impact of the pH and carbon source. Appl Microbiol Biotechnol 80:1121–1130. https://doi.org/10.1007/s00253-008-1669-x.

70. Roels J. 1983. Energetics and kinetics in biotechnology. Biomedical Press,Amsterdam, Netherlands.

71. van den Berg EM, Elisário MP, Kuenen JG, Kleerebezem R, van Loos-drecht MCM. 2017. Fermentative bacteria influence the competitionbetween denitrifiers and DNRA bacteria. Front Microbiol 8:1–13. https://doi.org/10.3389/fmicb.2017.01684.

72. Wang Y, Qian PY. 2009. Conservative fragments in bacterial 16S rRNAgenes and primer design for 16S ribosomal DNA amplicons in meta-genomic studies. PLoS One 4:e7401. https://doi.org/10.1371/journal.pone.0007401.

73. Johnson K, Jiang Y, Kleerebezem R, Muyzer G, van Loosdrecht MCM.2009. Enrichment of a mixed bacterial culture with a high polyhydroxy-

alkanoate storage capacity. Biomacromolecules 10:670 – 676. https://doi.org/10.1021/bm8013796.

74. Nurk S, Meleshko D, Korobeynikov A, Pevzner PA. 2017. MetaSPAdes: a newversatile metagenomic assembler. Genome Res 27:824–834. https://doi.org/10.1101/gr.213959.116.

75. Li H, Durbin R. 2010. Fast and accurate long-read alignment withBurrows-Wheeler transform. Bioinformatics 26:589 –595. https://doi.org/10.1093/bioinformatics/btp698.

76. Li H, Handsaker B, Wysoker A, Fennell T, Ruan J, Homer N, Marth G,Abecasis G, Durbin R. 2009. The Sequence Alignment/Map formatand SAMtools. Bioinformatics 25:2078 –2079. https://doi.org/10.1093/bioinformatics/btp352.

77. Sieber CMK, Probst AJ, Sharrar A, Thomas BC, Hess M, Tringe SG, BanfieldJF. 2017. Recovery of genomes from metagenomes via a dereplication,aggregation, and scoring strategy. Nat Microbiol 3:836 – 843. https://doi.org/10.1038/s41564-018-0171-1.

78. Kempf VAJ, Trebesius K, Autenrieth IB. 2000. Fluorescent in situ hybrid-ization allows rapid identification of microorganisms in blood cultures. JClin Microbiol 38:830 – 838.

79. Kong Y, He M, McAlister T, Seviour R, Forster R. 2010. Quantitativefluorescence in situ hybridization of microbial communities in the ru-mens of cattle fed different diets. Appl Environ Microbiol 76:6933– 6938.https://doi.org/10.1128/AEM.00217-10.

80. Amann RI, Binder BJ, Olson RJ, Chisholm SW, Devereux R, Stahl DA. 1990.Combination of 16S rRNA-targeted oligonucleotide probes with flowcytometry for analyzing mixed microbial populations. Appl EnvironMicrobiol 56:1919 –1925.

81. Daims H, Brühl A, Amann R, Schleifer K-H, Wagner M. 1999. Thedomain-specific probe EUB338 is insufficient for the detection of allbacteria: development and evaluation of a more comprehensiveprobe set. Syst Appl Microbiol 22:434 – 444. https://doi.org/10.1016/S0723-2020(99)80053-8.

Anaerobic Galacturonate Enrichment Cultures Applied and Environmental Microbiology

September 2018 Volume 84 Issue 18 e01370-18 aem.asm.org 15

on April 5, 2020 by guest

http://aem.asm

.org/D

ownloaded from