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Hydrogen-based syntrophy in an electrically conductive biofilm anode
Accepted Manuscript Hydrogen-based syntrophy in an electrically conductive biofilm anode Bipro Ranjan Dhar, Jeong-Hoon Park, Hee-Deung Park, Hyung-Sool Lee PII: DOI: Reference:
S1385-8947(18)32372-6 https://doi.org/10.1016/j.cej.2018.11.138 CEJ 20440
To appear in:
Chemical Engineering Journal
Received Date: Revised Date: Accepted Date:
30 September 2018 15 November 2018 18 November 2018
Please cite this article as: B.R. Dhar, J-H. Park, H-D. Park, H-S. Lee, Hydrogen-based syntrophy in an electrically conductive biofilm anode, Chemical Engineering Journal (2018), doi: https://doi.org/10.1016/j.cej.2018.11.138
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Hydrogen-based syntrophy in an electrically conductive biofilm anode Bipro Ranjan Dhara, Jeong-Hoon Parkb, Hee-Deung Parkb, Hyung-Sool Leec* a
Civil and Environmental Engineering, University of Alberta, 9211-116 Street NW, Edmonton, Alberta T6G 1H9, Canada b Department of Civil, Environmental and Architectural Engineering, Korea University, Seoul 02841, South Korea c
Civil & Environmental Engineering, University of Waterloo, 200 University Ave W, Waterloo ON, N2L 3G1, Canada
*Corresponding author: Phone: +1-519-888-4567 Ext. 31095; Fax: +1-519-888-4349; E-mail: [email protected] Abstract We experimentally and theoretically investigated implications of H2 and a rate-limiting step in a mixed-culture biofilm anode fed with n-butyrate, one of the poorest substrates to exoelectrogens. Acetate and i-butyrate were formed as intermediates during anaerobic degradation of n-butyrate, which suggested oxidative acetogenesis of n-butyrate in syntrophy with H2 scavengers in the biofilm anode. Methane was not detected in an anode chamber, and no current was generated in the biofilm anode using H2 as the electron donor. These results indicated that acetogens would be a main H2 consumer in the biofilm. Pyrosequencing data showed dominance of Geobacter in the biofilm anode (83.6% of total sequences), along with Sphaerochaeta and Treponema, which supports the syntrophy between exoelectrogens and acetogens. Electrical conductivity of the butyrate-fed biofilm anode was as high as 0.67 mS/cm, demonstrating that EET does not limit current density in the biofilm. In-situ monitoring of dissolved H2 concentration proved H2 production (up to 12.4 µM) and consumption during current generation in the biofilm, which supports significance of H2–based syntrophy in the electrically conductive biofilm using n-butyrate as the primary electron donor.
Keywords: butyrate, hydrogen, Geobacter, biofilm conductivity, acetogenesis, direct interspecies electron transfer
Abbreviations [H+], proton concentration (M); [HAc], acetate concentration (M); [HBu], total butyrate concentration (M) including n-butyrate and ibutyrate; EET, extracellular electron transfer; GBiofilm (obs), intrinsic biofilm conductance (mS); GBiofilm (obs), observed biofilm conductance (mS); GControl, ionic conductance (mS); L, length of the electrodes (cm); Lf, biofilm thickness (μm); MxC, microbial electrochemical cell; OUT, operational taxonomic unit; PH2, hydrogen partial pressure (atm); R, ideal gas constant (0.00831 kJ/molK); SHE, standard hydrogen electrode (V); T, temperature (K); ΔG', standard Gibbs free energy (kJ/mol)
1. Introduction Low current density has been commonly found in microbial electrochemical cells (MxCs) treating fermentable substrate, although producing high current density is essential for MxC application as sustainable wastewater treatment [1–5]. For instance, dark fermentation was integrated with MxCs to improve H2 yield, but H2 production rate (proportional to current density) in MxCs is several orders of magnitude slower than that in dark fermentation [1,6]. The poor kinetics of biofilm anodes to fermentable substrate is one of the main bottlenecks to MxC deployment in field [3,6,7]. To overcome this kinetic challenge, hydrolysis and fermentation should be well symphonised with anode respiration in biofilm anodes. Previous studies have suggested that exoelectrogens would utilize fermentation products (e.g., propionate, ethanol or butyrate) in syntrophy with fermenting bacteria and H2 consumers in mixed culture biofilm anodes [1,8–10], while some exoelectrogens are able to oxidize them directly [10,11]. By keeping low partial pressure of H2 (PH2) in the biofilms, H2 consumption can play a vital role in driving oxidative acetogenesis of fermentable organics (e.g., ethanol conversion to acetate and H2), which enables exoelectrogens to oxidize acetate for current generation in MxCs [1,9,12]. H2-oxidizing exoelectrogens, hydrogenotrophic methanogens, or acetogens can scavenge the H2 produced from oxidative acetogenesis to meet the requirement of a threshold H2 concentration below which the acetogenesis can occur thermodynamically. However, no studies have experimentally proved H2 consumption in biofilm anodes using fermentable substrate. Literature proposes the syntrophic interaction among fermenters, H2 scavengers and exoelectrogens using microbial community data targeting 16S-rRNA gene, together with chemical analysis of metabolites [9,12]; DNA-based approach cannot prove the proposed syntrophy, which means that the microbial syntrophy in the biofilm anodes is still hypothetical.
There is limited information on kinetics on mixed-culture biofilm anodes utilizing fermentable substrate despite of low current density commonly observed in the biofilms [1,9,13]. Intrinsic electron transfer from fermentable substrate to the anode can be simplified into two steps of intra/intercellular electron transfer and extracellular electron transfer (EET) [7,14,15]. Intra and intercellular electron transfer would consist of the complexity of microbial syntrophy including fermentation of primary electron donor, H2 consumption, and acetate oxidation. In comparison, EET solely accounts for the electron transfer from outer membrane proteins of exoelectrogens to an anode via biofilms. As compared to acetate-fed biofilms, neither intracellular electron transfer nor EET kinetic has been studied thoroughly for the biofilm fed with fermentable substrate. Literature has reported that acetate-fed biofilms are electrically conductive, showing high biofilm conductivity from 0.25 to ~ 5 mS/cm [16–19]. Lee et al. [18] proposed that intracellular electron transfer would mainly limit current density in acetate-fed conductive biofilms because EET kinetic could account for several hundreds A/m2. In comparison, no studies have explored the electrical conductivity of biofilm anodes fed with fermentable substrate, and thus we do not understand which step between intra/intracellular and extracellular electron transfer limits current density in the biofilm anodes. In this work, we comprehensively studied a biofilm anode fed with n-butyrate, one of the poorest electron donors to exoelectrogens to explore butyrate degradation pathway and a rate-limiting step to current production in the biofilm. The goal of this study is four-fold. The first is to demonstrate the requirement of syntrophic interactions among fermenters, H2 consumers, and exoelectrogens to improve current density in butyrate-fed MxCs. The second is to prove syntrophic interactions associated with threshold H2 concentration by multiple approaches of thermodynamic analysis, molecular biology and experimental measurements of
dissolved H2. The third is to prove butyrate degradation pathway, and the final is to quantify the electrical conductivity of the butyrate-fed biofilm anode to assess a limitation step for current density. To the best of our knowledge, this study first presents experimental demonstration on dissolved H2 concentration and biofilm conductivity for the biofilm anode fed with butyrate.
2. Materials and Methods 2.1 Microbial Electrochemical Cells (MxCs) Four dual-chamber MxCs (MxC-1, MxC-2, MxC-3, and MxC-4) were fabricated for this study (see Fig. 1). MxC-1 was constructed with plexiglass tubes as previously described in the literature [8] , and was equipped with carbon fibers (2293-A, 24A Carbon Fiber, Fibre Glast Development Corp., Ohio, USA) integrated with a stainless-steel frame as the anode module (Fig. 1(a)). MxC-2, a H-type MxC in Fig. 1(b), was constructed with glass bottles (250 mL for each chamber) and a cube-shaped carbon felt (3 cm 3 cm 3 cm) was used as the anode. Stainless steel mesh was used as the cathode for MxC-1, MxC-2, and MxC-4. MxC-2 was specifically designed for continuously monitoring dissolved H2 concentration near the anode surface, allowing evaluation of n-butyrate fermentation to acetate and H2 (oxidative acetogenesis). MxC-3, shown in Fig. 1(c), was designed for measuring the electrical conductivity of anode biofilms. Two gold electrodes separated by a non-conductive gap of 50 μm were used as the anodes and biofilm conductivity was measured once biofilms bridged the two anodes. The detailed description of the MxC-3 could be found in the literature [19]. Briefly describing, the dual-chamber MxC-3 was constructed with plexiglass. Two gold electrodes (width
9.5 mm × length 15 mm × thickness 10 μm; total surface are of 2.85 cm2) on a glass base with a non-conductive gap of 50 μm was used as the anode electrode; while a porous graphite plate (Isomolded Graphite Plate 203101, Fuel Cell Earth, USA) was used as the cathode electrode for convenient design and operation. A microscopic image of the non-conductive gap between two gold electrodes were provided in the Supplementary Information. The working volumes of both chambers were 15 mL. MxC-4 was identical to MxC-1 and was used for H2-sparging experiment to assess whether H2 can be directly utilized as an electron donor. We employed anion exchange membranes (AMI-7001, Membranes International Inc., USA) as separators for the four MxCs to keep neutral pH in anolytes. All the MxCs were operated as three-electrode mode (anode, cathode and reference electrode) with an Ag/AgCl reference electrode (MF-2052, Bioanalytical System Inc., USA) placed ~1 cm apart from anodes. During operation of the four MxCs, the anode potential was constantly set at -0.2 V vs. SHE using a multi-channel potentiostat system (BioLogic, VSP, Gamble Technologies, Canada). Currents were recorded at every 120 s using EC-Lab for Windows v10.12 software in a personal computer connected to the potentiostat. 2.2 Enrichment of Anode Biofilms and Operation of MxCs Initially, the anode biofilm in MxC-1 was inoculated with 20 mL of effluent from a mother MxC that had been run with 25 mM acetate medium in over 1.5 years; Geobacter was highly enriched at 98% in the biofilm anode of the mother MxC [19]. To enrich butyrate-fermenting bacteria and H2 consumers (e.g., acetogens or methanogens) we added 20 mL of anaerobic sludge collected from an anaerobic digester at St Mary’s wastewater treatment plant (Ontario, Canada). Then, we filled the anode chamber with butyrate
medium (~12 mM n-butyrate equivalent to 1,954±156 mg COD/L) supplemented with 50 mM phosphate buffer. The literature provides detailed information on mineral composition [8]. The cathode chamber was filled with tap water where H2 gas was produced. We operated MxC-1 in fed-batch mode and replaced anolyte with fresh butyrate medium after the current density dropped below 0.5 A/m2 for each cycle. We kept operating fed-batch operation of MxC-1 until current change to time was replicated (~1.5 months), and after that data was collected for analysis. The anode biofilms collected from the MxC-1 with a sterilized spatula were transferred to the other three MxCs as inocula. To avoid oxygen exposure to exoelectrogens, collection and inoculation of biofilms were conducted in an anaerobic chamber (COY Type B Vinyl Anaerobic Chamber, COY Lab Products, USA). MxC-2 was consistently run in batch mode using the butyrate medium to accurately measure dissolved H2 concentration with time in an anode chamber. MxC-3 were initially operated with butyrate medium in batch mode until producing positive current density. Then, we switched MxC-3 into continuous mode by feeding butyrate medium with a cartridge-type peristaltic pump (Master Flex® L/S digital drive, Model 7523-80, Cole-Parmer, Canada) to maintain a hydraulic residence time (HRT) of 2.6 h. After the steady-state current density reached at ~1.15 A/m2 in MxC-3, the electrical conductivity of the biofilm anode was measured. MxC-4 was operated using H2 as the electron donor to confirm if exoelectrogens prefer to consume H2 for current generation; we summarized inoculation and operation of MxC-4 in the Supplementary Material. All experiments were conducted in a temperature controlled room at 25 ± 1°C. 2.3. Serum bottle tests To assess butyrate fermentation without anode respiration of exoelectrogens, biofilms and supernatant 5 mL collected from MxC-1 were transferred to serum bottles having 10 mL of the butyrate medium; the total volume of serum bottle was 20 mL (working volume
15 mL and headspace 5 mL). After sparging the bottles with ultra-pure nitrogen (99.999%) for 1 min, the liquids in the serum bottles were continuously mixed at 100 rpm using a magnetic stirrer (Model 650, VWR International Inc., Canada) in the temperaturecontrolled room. We measured initial and final concentrations of volatile fatty acids in liquid samples. The volume and composition of biogas produced from serum bottles were intermittently monitored. The total duration of the serum bottle test was 72 hours. Serum bottle tests were conducted in duplicates, and average data was used for calculating cumulative H2 production, according to the literature [13].
2.4 Analytical Methods We quantified volatile fatty acids using a gas chromatography (GC) (Model: Hewlett Packard HP 5890 Series II) equipped with a Nukol fused-silica capillary column and flame ionization detector (FID) using He as a carrier gas. The initial temperature of the column was 110°C, increasing to 195°C at the rate of 8°C/min, and then held constant at the final temperature of 195°C for 9 min. Injector and detector temperatures were maintained at 220°C and 280°C, respectively. The liquid samples were acidified to pH ~ 2 using 1 N phosphoric acid prior to GC-FID analyses, and then filtered using 0.2 µm membrane syringe filter. As confirmed in our previous study [9], the detection limits for acetate, n-butyrate, and i-butyrate were 0.03, 0.02, 0.004 mM, respectively.
All liquid
samples were analyzed in triplicates. The volume of biogas produced in serum bottle tests was measured with a friction-free glass syringe (Perfektum Glass Luer Lock Syringe 30 mL, Thomas Scientific, NJ, USA). H2 percentage in the biogas was quantified with a
GC (Model 310, SRI Instruments, Torrance, CA) equipped with a thermal conductivity detector (TCD) and a molecular sieve column (Molesieve 5A, mesh 80/100, 6 ft × 1/8 in.). Argon (99.999%, PraxAir, Canada) was used as a carrier gas with a flow rate of 10 mL/min under a pressure of 21 psi for the GC. The temperatures of the column oven and the detector of the GC were kept at 31°C and 200°C, respectively. The dissolved H2 concentration in MxC-1 and MxC-2 was measured using a H2 microelectrode (Unisense H2-100, Unisense A/S, Denmark) with a tip diameter of 100 m connected with a 4-channel microsensor multimeter (Microsensor Multimeter for Unisense Sensors, 2x pA, 1x mV and 1x Temp channel, Unisense A/S, Denmark). Using 20 mL vials (10 mL deionized water and 10 mL headspace) we prepared different H2 concentrations in liquid and H2 gas compositions in headspace by sparging the vials with H2 and N2 gases (99.999%, PraxAir, Canada) in given times. The vials were vigorously shaken with a vortex mixer for 3 minutes at 3,000 µ at room temperature. The percentage of H2 gas in the headspace was measured with the GC-TCD and converted to the concentration of dissolved H2 in liquid using Henry’s law constant for H2 (0.00078 mol/L-atm at 25°C), giving a range of 13-399 µM of aqueous H2. We then built a calibration curve between the prepared H2 concentrations and the voltage values read by the H2 microelectrode (R2=0.99). We measured the dissolved H2 concentration in the anolyte of MxC-1 by inserting the H2 microsensor through a gas-tight rubber on top of MxC-1; the microsensor was 7 cm distant from the anode in MxC-1. H2 molecules can be consumed by exoelectrogens, methanogens or acetogens in biofilm anodes or bulk liquid, which can bias aqueous H2 measurements with the microsensor. To overcome this challenge, we used MxC-2 and positioned the H2 microelectrode tip in the vicinity of the anode using
a motorized micromanipulator (Unisense MM33, Unisense A/S, Denmark). We then measured dissolved H2 concentration over time using SensorTrace Suite v2.1.100 software in a computer connected to the microsensor multimeter. The electrical conductivity of the anode biofilms was measured using MxC-3 with the two-probe method according to the literature [18–20]. Briefly describing, the gold anodes and the cathode were disconnected temporarily (open circuit mode). Then, a linear sweeping voltage of 0-0.05 V in a step of 0.025 V was applied across two gold electrodes using a source measurement unit (Keithley 2400, Keithley Instruments, Inc., USA), and the current was recorded for each voltage. The voltage ramp was applied in 4-5 cycles until a steady-state current-voltage (I-V) response was obtained. Observed biofilm conductance (GBiofilm (obs), mS) was calculated from steady-state I-V curves. The ionic conductance (Gcontrol, mS) was measured with an abiotic MxC-3 to account for ionic current across the non-conductive gap using the butyrate medium. Intrinsic biofilm conductance was determined by considering ionioc conductance (GBiofilm = GBiofilm (obs) - Gcontrol, mS) and we calculated biofilm conductivity with Eq. (1) [21].
Where, Gbiofilm is intrinsic biofilm conductance (mS) (GBiofilm = GBiofilm (obs)-Gcontrol), Lf is the biofilm thickness (μm), L is the length of the electrodes (1.5 cm), and a is half of the non-conductive gap between two electrodes (25 μm). Biofilm thickness in MxC-3 was measured with a microelectrode as previously described in the literature [14,19]. Supplementary Material provided details of biofilm thickness measurement.
2.5 Microbial Community Analysis We collected biofilm and suspended cells in MxC-1 to analyze microbial community. No methane was detected in this study, and hence we focused on bacterial community analysis. At the end of MxC-1 operation (days 120), we disintegrated MxC-1 in the anaerobic chamber and took a portion of biofilms with a sterilized spatula. We transferred the anolyte into multiple microcentrifuge tubes (~1.5 mL in each tube) and centrifuged them at 5,000 rpm for 2 min (Eppendorf 5424, Hamburg, Germany) to collect pellets. The biofilm and planktonic cells were stored at -20 °C before DNA extraction and sequencing steps. DNA was extracted using the MoBio PowerSoil DNA extraction kit (Solana Beach, CA, USA) according to the manufacturer’s provided protocol. Concentration and purity of DNA were measured using the NanoDrop ND-1000 (NanoDrop Technologies, Wilmington, DE, USA). Typically the values were around 50 ng/μL and around 1.9 (Abs 260 nm/Abs280 nm), respectively. Barcode 454 pyrosequencing targeting bacterial 16S rRNA genes was conducted using the extracted community DNA at the Macrogen (Seoul, South Korea). For the amplification of bacterial 16S rRNA genes, bacterial universal primers (27F: 5´-AGAGTTTGATCMTGGCTCAG-3´, 518R: (5´ATTACCGCGGCTGCTGG-3´) were used [22,23]. The detailed procedure of pyrosequencing was described in a previous study [24]. From sequencing results, species richness (Chao1 and ACE indices) and diversity (Shannon and Simpson indices) were calculated using the Mothur utility [24]. Operational taxonomic units (OTUs) were defined as groups of sequences showing > 97% sequence identities using the Mothur utility. Taxonomic identities of the OTUs were determined based on the SILVA database (https://www.arb-silva.de).
3. Results and Discussions 3.1 Butyrate consumption in a biofilm anode Fig. 2-(a) shows the evolution of current density with time in two consecutive batch operations of MxC-1 after we confirmed four replicate patterns of current density to time. For both runs, the current density rapidly peaked at 7.8-8.8 A/m2 after addition of fresh butyrate medium, and then decreased in a week approximately. Like current density, the concentration of n-butyrate gradually reduced with time, as shown in Fig. 2-(b). i-butyrate (0.17-4.9 mM) and acetate (0.35-14.7 mM) were intermediately accumulated and degraded with time. Isomerization of n-butyrate was commonly found as an intermediate in n-butyrate degradation of methanogenic enrichments [25,26]. The intermittent accumulation of acetate and i-butyrate from n-butyrate suggests that butyrate fermentation to acetate and H2 (oxidative acetogenesis) would occur in MxC-1 (C4H7O2- + 2H2O → 2CH3COO- + H+ + 2H2). This fermentation reaction, however, is thermodynamically unfavorable under standard conditions at pH 7; the standard Gibbs free energy at pH 7 (ΔG') for the n-butyrate fermentation is +48.3 kJ/mol butyrate [1,27]. Aqueous H2 concentration (or partial pressure of H2 (PH2)) should be maintained very low for the butyrate fermentation to occur. Hence, the degradation of n-butyrate and intermittent accumulation of ibutyrate and acetate imply that H2-oxidizing microorganisms would consume H2 produced from the butyrate fermentation, overcoming the thermodynamic bottleneck in the oxidative acetogenesis of butyrate. This interpretation well accords to the literatures [1,9,28], suggesting syntrophic interactions among fermenters, H2 consumers and exoelectrogens in biofilm anodes fed with fermentable substrates.
3.2 Thermodynamic evaluation of butyrate fermentation in the biofilm anode: threshold H2 concentration To drive the butyrate fermentation in MxC-1, dissolved H2 concentration should be less than threshold H2 concentration that is able to trigger butyrate fermentation to acetate and H2 thermodynamically. According to Eq. (2) we computed the threshold PH2 and converted it to the threshold H2 concentration using Henry’s law constant for H2 (0.00078 mol/L-atm at 25°C).
Where, ΔG' is the standard Gibbs free energy for butyrate fermentation (+48.3 kJ/mol butyrate), R is the ideal gas constant (0.00831 kJ/mol-K), T is the temperature (K), [HBu] is the total butyrate concentration (M) including n-butyrate and i-butyrate, [HAc] is the acetate concentration (M), and [H+] is the proton concentration (M). For thermodynamic calculation, we assumed that i-butyrate is transformed to n-butyrate and the n-butyrate is fermented to acetate and H2 [26,29]; energy change in isomerization of n-butyrate is negligible [25], so we ignored small energy change in butyrate isomerization for this computation. Fig. 3 shows the evolution of the threshold H2 concentration in MxC-1 when n-butyrate, i-butyrate, and acetate concentration were varied with time (Fig. 2). Thermodynamic evaluation on the butyrate fermentation indicates that the threshold H2 concentration should range from 0.1 to 7.2 μM (PH2 10-942 Pa), indicating that H2 gas should not be accumulated in the anodic headspace of MxC-1. We did not detect any H2 gas in the headspace with the GC-TCD (detection limit > 0.01 atm H2), and this result accords to the threshold H2 calculation in Fig. 3. We measured the dissolved H2 concentration in the anolyte of MxC-1 with the H2 microsensor that
can precisely quantify a few μM ranges of dissolved H2 with a detection limit of 0.3 µM (www.unisense.com). Unexpectedly, no dissolved H2 was detected in the anolyte, which contradicts to the thermodynamic analysis of threshold H2 computation. The microsensor was located at 7 cm distant from the anode in MxC-1. H2 molecules produced from the butyrate fermentation could be scavenged by H2 consumers in the biofilm anode or bulk liquid before the H2 reaches the microsensor; in Section 3.4 we further explored dissolved H2 concentration in the butyrate fermentation using MxC-2 where the microsensor physically touched the anode surface to mitigate H2 consumption effect during measurements. Hydrogenotrophic methanogens, H2-oxidizing exoelectrogens, or acetogens can consume H2 produced from butyrate fermentation. We did not detect any methane gas in the headspace of MxC-1 during experiments, excluding the methanogens from H2 consumer candidates. To test H2 oxidation by exoelectrogens, we operated MxC-4 using H2 gas as the sole electron donor, but current density in the H2-fed MxC was negligible at 0.03-0.24 A/m2 close to endogenous decay current density (see Supplementary Material for details). Thus, direct H2 oxidation by exoelectrogens would be negligible for H2 consumption in MxC-1. It is likely that acetogens would be main H2 consumers in MxC-1, suggesting significance of the syntrophy between acetogens and exoelectrogens for current generation (H2 + CO2 → acetate → electrical current). Several studies proposed this syntrophic interaction in MxCs fed with fermentable substrate [1,9,12]. In serum bottle test, the concentration of n-butyrate slightly decreased in 72 h, along with small increase of i-butyrate and acetate, as described in Table 1. Cumulative H2 production was small at 0.14±0.03 mL H2 and its production was stopped at 48 h. No methane was detected during the experiments, consistent to MxC-1, and the concentration of the acids was not changed after 72 h of reaction
time (data not shown). The evolution of VFAs and H2 in serum bottle tests supports oxidative acetogenesis of n-butyrate (butyrate conversion to acetate and H2) in MxC-1, but the cease of H2 production and n-butyrate degradation proves significance of exoelectrogens and their syntrophy with fermenters and H2 consumers for continuing the oxidative acetogenesis.
3.3 Microbial community in biofilm and planktonic cells From the two samples (biofilm and suspended cells in MxC-1) 1,036 – 1,450 of sequences were used for bacterial community analysis after removal of chimera and short sequences (< 250 base pairs). Table S1 compares species (OTU) richness and diversity estimated from the retrieved 16S rRNA gene sequences. Both two species richness indices (Chao1 and ACE) estimated lower richness of bacteria species in the biofilm than planktonic cells. The same pattern was also observed in two species diversity indices (Shannon and Simpson). Bacterial community compositions were compared in Fig. 4. Communities were not significantly different at the phylum level (see Fig. 4(a)). The most dominant phylum was Proteobacteria for the biofilm and planktonic cells (61.3 – 90.5%), following Firmicutes, Synergistetes, Bacteroidetes, and Spirochaetes. However, communities were very different at the genus level, as shown in Fig. 4(b). Geobacter was predominant (83.6% out of total sequences) in the biofilm anode, like the mother MxC. Small populations of Sphaerochaeta and Treponema, that can build the syntrophy with butyrate fermenters as acetogens [30,31], were identified along
with 11.85% unclassified bacteria. This result indicates that Geobacter is the main exoelectrogen in the biofilm anode, as reported by the literature [9,18–20,32]. The biofilm community supports the proposed syntrophy between exoelectrogens and acetogens. In comparison, Geobacter only comprised 2.1% and the other genera were dominant in planktonic cells: Pseudomonas (18.7%), Aeromonas (16.4%), Sphaerochaeta (8.2%), Treponema (3.8%), and Proteiniphilum (3.5%). Metabolic functions of Pseudomonas and Aeromonas are versatile [33,34], and hence it is challenging to assess their metabolic functions with 16S rRNA genes. Pseudomonas and Aeromonas can work as fermenters [35,36] or conduct anaerobic respiration using conductive solids [37,38]. For instance, some Pseudomonas species can produce electron shuttling compounds (e.g. pyocyanin) for mediated electron transport to solid conductors [38,39]. Both Pseudomonas and Aeromonas species also possess genes for EET and can produce the electrical current in MxCs [37,40,41]. Consistent to the biofilm, Sphaerochaeta and Treponema were also identified in the suspended cells, acetogen candidates. The bacterial community of the planktonic cells implies that different exoelectrogens might play a role of acetate consumption in bulk liquid, but consistently supports the proposed syntrophy of acetogens and exoelectrogens. 3.4 H2 production and consumption in the butyrate-fed biofilm anode Fig. 5 shows the change of current density and dissolved H2 concentration with time in MxC-2. Like the batch run of MxC-1, current density gradually increased, reached a plateau, and decreased with time. Dissolved H2 concentration was as high as 12.4 µM (PH2 1,640 Pa) at the initial fermentation of n-butyrate, which indicates H2 production in oxidative acetogenesis of butyrate. Dissolved H2 concentration decreased with time, and it became as small as 3.5 µM at the end of operation (140 h). This result clearly evidences H2 consumption in MxC-2; here, we only sampled and analyzed liquid samples at the end of batch run because sampling readily disturbed
the measurements of dissolved H2 concentration with the microsensor. The measured H2 concentration was very close to 3.1 µM of threshold H2 concentration calculated with Eq. 2 (n-butyrate 2.6 mM, i-butyrate 1.3 mM, and acetate 0.9 mM), validating the thermodynamic evaluation of butyrate fermentation and threshold H2 calculation discussed early in Section 3.2. Abrupt increase of current density involved sharp decrease of dissolved H2 concentration in the biofilm anode (an area highlighted with blue color in Fig. 5). This experimental result first proves the significance of H2 consumption for exoelectrogens to generate electric current in the butyrate-fed biofilm anode. Based on molecular biology data, the syntrophic interaction between exoelectrogens and acetogens is a key to overcome a thermodynamic barrier in utilization of butyrate in the biofilm anode. 3.5 Biofilm conductivity Fig. 6 shows the biofilm conductivity measured with MxC-3. The biofilm conductivity was relatively stable at 0.67±0.14 mS/cm in 12 measurements for consecutive four days (biofilm thickness 138±19 μm). The relatively large standard deviation in the average biofilm conductivity would indicate the requirement of a more robust method on biofilm conductivity. However, this biofilm conductivity is within the range of biofilm conductivities (0.25-5 mS/cm) observed for Geobacter-enriched biofilm anodes [14,16,17,19]. This experimental result first proves that the butyrate-fed biofilm is electrically conductive, although complex syntrophic interactions among fermenters, H2 consumers and exoelectrogens can be built in the biofilm.
This high biofilm
conductivity means that electrical conduction governs EET to the anode in the biofilm. According to Eq. (3), the maximum current density achievable in the EET step is computed at ~ 30 A/m2. This calculated current density is even several times higher than the highest current density in acetate-fed biofilm anodes [18,42,43].
j = Kbio (Eanode – EEC° )/Lf
(3)
Where, Kbio is biofilm conductivity (S/m), Eanode is anode potential (V) (-0.2 V vs. SHE), and EEC° is the standard potential for a ratelimiting extracellular cofactor at pH 7 (V). We assumed EEC° = -0.26 V vs. SHE for calculating the maximum current density [18,44]. Low current density (< 2 A/m2) has been commonly observed in MxCs using butyrate as substrate [45,46], but we have not well understood which step causes such low current density in butyrate-fed MxCs. Our study indicates that intra and intercellular electron transfer would mainly limit current density in butyrate-fed biofilms, not EET. The kinetic assessment of intracellular and intercellular electron transfer in the butyrate-fed biofilm is not straightforward, due to complex microbial syntrophy including butyrate fermentation, acetate oxidation, H2 consumption, and mass transfer of intermediate metabolites (e.g., H2 and acetate transfer). More study is required to investigate a rate-limiting step in the intracellular and intercellular electron transfer kinetics for improving current density in butyrate-fed MxCs. The electrically conductive biofilm implies that direct interspecies electron transfer (DIET) might occur among butyrate fermenters, H2 consumers, and exoelectrogens, as proposed by recent studies [47,48]. DIET is an efficient manner to build electronsharing networks among microorganisms [48–51], but our study proved the significance of H2 syntrophy in the electrically conductive biofilm anode. For inter-electron transport, DIET may be better than hydrogen molecules considering mass transport limitation of H2.
In energetic point of view, however, H2 syntrophy would be superior to DIET because H2 is a great energy-carrier to syntrophic partners; in comparison, no studies have proved energy benefit to microorganisms in DIET.
4. Conclusions A high electrical conductivity (0.67±0.14 mS/cm) was measured for the butyrate-fed biofilm, which demonstrates that EET does not limit current density in the biofilm. Despite of high electrical conductivity in the biofilm, in-situ measurements of H2 concentration confirmed H2 production and consumption consistent to current generation in the butyrate-fed biofilm. Geobacter was identified as the main exoelectrogen in syntrophy with Sphaerochaeta and Treponema, supporting the significant roles of acetogens as the H2 scavenger and syntrophic partner in the biofilm. The high electrical conductivity of the biofilm suggests that DIET might occur to link exoelectrogens with syntrophic partners for butyrate utilization, but this study proves the significance of H2-based syntrophy even in the electrically conductive biofilm. Acknowledgements This research was funded by Natural Sciences and Engineering Research Council of Canada Discovery Grant (RGPIN-2016-04155). We appreciate Dr. Jangho Lee’s supports to calibration of the H2 microsensor. References
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Table and figure captions Table 1. Changes of acids and biogases in serum bottles lacking anode respiration.
Fig. 1. Schematic diagram showing configuration of (a) MxC-1 and MxC-4, (b) MxC-2, and (c) MxC-3. Note. MxC-2 and MxC-3 were designed for quantifying dissolved hydrogen concentration and biofilm conductivity during butyrate fermentation, respectively. Figures are not drawn to scale. Fig. 2. Profiles of current density and main VFAs to time in MxC-1. (a) Current density, and (b) main VFAs of n-butyrate, i-butyrate, and acetate.
Fig. 3. The threshold H2 concentration for butyrate fermentation to acetate and H2 in MxC-1. The H2 concentration was computed with Eq. (2). Fig. 4. Relative abundance at (a) phylum, and (b) genus level. Fig. 5. Current density and dissolved hydrogen concentrations on anode surface in MxC-2 during fed-batch operation. Blue highlighted areas indicate abrupt increase of current density and decrease of dissolved H2 concentration in MxC-2. Fig. 6. Current density and measured biofilm conductivity during operation of MxC-3.
Table 1. Changes of acids and biogases in serum bottles lacking anode respiration.
Initial
Final
n-butyrate
10.8±0.1 mM
8.2±1.8 mM
i-butyrate
0.21±0.1 mM
0.8±0.2 mM
Acetate
0.6±0.3 mM
1.7±1.0 mM
H2 gas
0
0.14±0.03 mL
CH4 gas
0
N.D.
Serum bottle tests were conducted in 72 h. N.D.: not detected.
Highlights
The electrical conductivity of the butyrate-fed biofilm was as high as 0.67 mS/cm H2-based syntrophy played an important role in the electrically conductive biofilm Aqueous H2 concentration decreased as current increased Acetogens (Sphaerochaeta and Treponema) were primary H2 consumers in the biofilm Intracellular electron transport mainly limited current in the biofilm
Fig. 1. Schematic diagram showing configuration of (a) MxC-1 and MxC-4, (b) MxC-2, and (c) MxC-3. Note. MxC-2 and MxC-3 were designed for quantifying dissolved hydrogen concentration and biofilm conductivity during butyrate fermentation, respectively. Figures are not drawn to scale.
Fig. 2. Profiles of current density and main VFAs to time in MxC-1. (a) Current density, and (b) main VFAs of n-butyrate, i-butyrate, and acetate.
34
Fig. 3. The threshold H2 concentration for butyrate fermentation to acetate and H2 in MxC-1. The H2 concentration was computed with Eq. (2).
35
Fig. 4. Relative abundance at (a) phylum, and (b) genus level.
36
Fig. 5. Current density and dissolved hydrogen concentrations on anode surface in MxC-2 during fed-batch operation. Blue highlighted areas indicate abrupt increase of current density and decrease of dissolved H2 concentration in MxC-2.
37
Fig. 6. Current density and measured biofilm conductivity during operation of MxC-3.
38
S1385-8947(18)32372-6 https://doi.org/10.1016/j.cej.2018.11.138 CEJ 20440
To appear in:
Chemical Engineering Journal
Received Date: Revised Date: Accepted Date:
30 September 2018 15 November 2018 18 November 2018
Please cite this article as: B.R. Dhar, J-H. Park, H-D. Park, H-S. Lee, Hydrogen-based syntrophy in an electrically conductive biofilm anode, Chemical Engineering Journal (2018), doi: https://doi.org/10.1016/j.cej.2018.11.138
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Hydrogen-based syntrophy in an electrically conductive biofilm anode Bipro Ranjan Dhara, Jeong-Hoon Parkb, Hee-Deung Parkb, Hyung-Sool Leec* a
Civil and Environmental Engineering, University of Alberta, 9211-116 Street NW, Edmonton, Alberta T6G 1H9, Canada b Department of Civil, Environmental and Architectural Engineering, Korea University, Seoul 02841, South Korea c
Civil & Environmental Engineering, University of Waterloo, 200 University Ave W, Waterloo ON, N2L 3G1, Canada
*Corresponding author: Phone: +1-519-888-4567 Ext. 31095; Fax: +1-519-888-4349; E-mail: [email protected] Abstract We experimentally and theoretically investigated implications of H2 and a rate-limiting step in a mixed-culture biofilm anode fed with n-butyrate, one of the poorest substrates to exoelectrogens. Acetate and i-butyrate were formed as intermediates during anaerobic degradation of n-butyrate, which suggested oxidative acetogenesis of n-butyrate in syntrophy with H2 scavengers in the biofilm anode. Methane was not detected in an anode chamber, and no current was generated in the biofilm anode using H2 as the electron donor. These results indicated that acetogens would be a main H2 consumer in the biofilm. Pyrosequencing data showed dominance of Geobacter in the biofilm anode (83.6% of total sequences), along with Sphaerochaeta and Treponema, which supports the syntrophy between exoelectrogens and acetogens. Electrical conductivity of the butyrate-fed biofilm anode was as high as 0.67 mS/cm, demonstrating that EET does not limit current density in the biofilm. In-situ monitoring of dissolved H2 concentration proved H2 production (up to 12.4 µM) and consumption during current generation in the biofilm, which supports significance of H2–based syntrophy in the electrically conductive biofilm using n-butyrate as the primary electron donor.
Keywords: butyrate, hydrogen, Geobacter, biofilm conductivity, acetogenesis, direct interspecies electron transfer
Abbreviations [H+], proton concentration (M); [HAc], acetate concentration (M); [HBu], total butyrate concentration (M) including n-butyrate and ibutyrate; EET, extracellular electron transfer; GBiofilm (obs), intrinsic biofilm conductance (mS); GBiofilm (obs), observed biofilm conductance (mS); GControl, ionic conductance (mS); L, length of the electrodes (cm); Lf, biofilm thickness (μm); MxC, microbial electrochemical cell; OUT, operational taxonomic unit; PH2, hydrogen partial pressure (atm); R, ideal gas constant (0.00831 kJ/molK); SHE, standard hydrogen electrode (V); T, temperature (K); ΔG', standard Gibbs free energy (kJ/mol)
1. Introduction Low current density has been commonly found in microbial electrochemical cells (MxCs) treating fermentable substrate, although producing high current density is essential for MxC application as sustainable wastewater treatment [1–5]. For instance, dark fermentation was integrated with MxCs to improve H2 yield, but H2 production rate (proportional to current density) in MxCs is several orders of magnitude slower than that in dark fermentation [1,6]. The poor kinetics of biofilm anodes to fermentable substrate is one of the main bottlenecks to MxC deployment in field [3,6,7]. To overcome this kinetic challenge, hydrolysis and fermentation should be well symphonised with anode respiration in biofilm anodes. Previous studies have suggested that exoelectrogens would utilize fermentation products (e.g., propionate, ethanol or butyrate) in syntrophy with fermenting bacteria and H2 consumers in mixed culture biofilm anodes [1,8–10], while some exoelectrogens are able to oxidize them directly [10,11]. By keeping low partial pressure of H2 (PH2) in the biofilms, H2 consumption can play a vital role in driving oxidative acetogenesis of fermentable organics (e.g., ethanol conversion to acetate and H2), which enables exoelectrogens to oxidize acetate for current generation in MxCs [1,9,12]. H2-oxidizing exoelectrogens, hydrogenotrophic methanogens, or acetogens can scavenge the H2 produced from oxidative acetogenesis to meet the requirement of a threshold H2 concentration below which the acetogenesis can occur thermodynamically. However, no studies have experimentally proved H2 consumption in biofilm anodes using fermentable substrate. Literature proposes the syntrophic interaction among fermenters, H2 scavengers and exoelectrogens using microbial community data targeting 16S-rRNA gene, together with chemical analysis of metabolites [9,12]; DNA-based approach cannot prove the proposed syntrophy, which means that the microbial syntrophy in the biofilm anodes is still hypothetical.
There is limited information on kinetics on mixed-culture biofilm anodes utilizing fermentable substrate despite of low current density commonly observed in the biofilms [1,9,13]. Intrinsic electron transfer from fermentable substrate to the anode can be simplified into two steps of intra/intercellular electron transfer and extracellular electron transfer (EET) [7,14,15]. Intra and intercellular electron transfer would consist of the complexity of microbial syntrophy including fermentation of primary electron donor, H2 consumption, and acetate oxidation. In comparison, EET solely accounts for the electron transfer from outer membrane proteins of exoelectrogens to an anode via biofilms. As compared to acetate-fed biofilms, neither intracellular electron transfer nor EET kinetic has been studied thoroughly for the biofilm fed with fermentable substrate. Literature has reported that acetate-fed biofilms are electrically conductive, showing high biofilm conductivity from 0.25 to ~ 5 mS/cm [16–19]. Lee et al. [18] proposed that intracellular electron transfer would mainly limit current density in acetate-fed conductive biofilms because EET kinetic could account for several hundreds A/m2. In comparison, no studies have explored the electrical conductivity of biofilm anodes fed with fermentable substrate, and thus we do not understand which step between intra/intracellular and extracellular electron transfer limits current density in the biofilm anodes. In this work, we comprehensively studied a biofilm anode fed with n-butyrate, one of the poorest electron donors to exoelectrogens to explore butyrate degradation pathway and a rate-limiting step to current production in the biofilm. The goal of this study is four-fold. The first is to demonstrate the requirement of syntrophic interactions among fermenters, H2 consumers, and exoelectrogens to improve current density in butyrate-fed MxCs. The second is to prove syntrophic interactions associated with threshold H2 concentration by multiple approaches of thermodynamic analysis, molecular biology and experimental measurements of
dissolved H2. The third is to prove butyrate degradation pathway, and the final is to quantify the electrical conductivity of the butyrate-fed biofilm anode to assess a limitation step for current density. To the best of our knowledge, this study first presents experimental demonstration on dissolved H2 concentration and biofilm conductivity for the biofilm anode fed with butyrate.
2. Materials and Methods 2.1 Microbial Electrochemical Cells (MxCs) Four dual-chamber MxCs (MxC-1, MxC-2, MxC-3, and MxC-4) were fabricated for this study (see Fig. 1). MxC-1 was constructed with plexiglass tubes as previously described in the literature [8] , and was equipped with carbon fibers (2293-A, 24A Carbon Fiber, Fibre Glast Development Corp., Ohio, USA) integrated with a stainless-steel frame as the anode module (Fig. 1(a)). MxC-2, a H-type MxC in Fig. 1(b), was constructed with glass bottles (250 mL for each chamber) and a cube-shaped carbon felt (3 cm 3 cm 3 cm) was used as the anode. Stainless steel mesh was used as the cathode for MxC-1, MxC-2, and MxC-4. MxC-2 was specifically designed for continuously monitoring dissolved H2 concentration near the anode surface, allowing evaluation of n-butyrate fermentation to acetate and H2 (oxidative acetogenesis). MxC-3, shown in Fig. 1(c), was designed for measuring the electrical conductivity of anode biofilms. Two gold electrodes separated by a non-conductive gap of 50 μm were used as the anodes and biofilm conductivity was measured once biofilms bridged the two anodes. The detailed description of the MxC-3 could be found in the literature [19]. Briefly describing, the dual-chamber MxC-3 was constructed with plexiglass. Two gold electrodes (width
9.5 mm × length 15 mm × thickness 10 μm; total surface are of 2.85 cm2) on a glass base with a non-conductive gap of 50 μm was used as the anode electrode; while a porous graphite plate (Isomolded Graphite Plate 203101, Fuel Cell Earth, USA) was used as the cathode electrode for convenient design and operation. A microscopic image of the non-conductive gap between two gold electrodes were provided in the Supplementary Information. The working volumes of both chambers were 15 mL. MxC-4 was identical to MxC-1 and was used for H2-sparging experiment to assess whether H2 can be directly utilized as an electron donor. We employed anion exchange membranes (AMI-7001, Membranes International Inc., USA) as separators for the four MxCs to keep neutral pH in anolytes. All the MxCs were operated as three-electrode mode (anode, cathode and reference electrode) with an Ag/AgCl reference electrode (MF-2052, Bioanalytical System Inc., USA) placed ~1 cm apart from anodes. During operation of the four MxCs, the anode potential was constantly set at -0.2 V vs. SHE using a multi-channel potentiostat system (BioLogic, VSP, Gamble Technologies, Canada). Currents were recorded at every 120 s using EC-Lab for Windows v10.12 software in a personal computer connected to the potentiostat. 2.2 Enrichment of Anode Biofilms and Operation of MxCs Initially, the anode biofilm in MxC-1 was inoculated with 20 mL of effluent from a mother MxC that had been run with 25 mM acetate medium in over 1.5 years; Geobacter was highly enriched at 98% in the biofilm anode of the mother MxC [19]. To enrich butyrate-fermenting bacteria and H2 consumers (e.g., acetogens or methanogens) we added 20 mL of anaerobic sludge collected from an anaerobic digester at St Mary’s wastewater treatment plant (Ontario, Canada). Then, we filled the anode chamber with butyrate
medium (~12 mM n-butyrate equivalent to 1,954±156 mg COD/L) supplemented with 50 mM phosphate buffer. The literature provides detailed information on mineral composition [8]. The cathode chamber was filled with tap water where H2 gas was produced. We operated MxC-1 in fed-batch mode and replaced anolyte with fresh butyrate medium after the current density dropped below 0.5 A/m2 for each cycle. We kept operating fed-batch operation of MxC-1 until current change to time was replicated (~1.5 months), and after that data was collected for analysis. The anode biofilms collected from the MxC-1 with a sterilized spatula were transferred to the other three MxCs as inocula. To avoid oxygen exposure to exoelectrogens, collection and inoculation of biofilms were conducted in an anaerobic chamber (COY Type B Vinyl Anaerobic Chamber, COY Lab Products, USA). MxC-2 was consistently run in batch mode using the butyrate medium to accurately measure dissolved H2 concentration with time in an anode chamber. MxC-3 were initially operated with butyrate medium in batch mode until producing positive current density. Then, we switched MxC-3 into continuous mode by feeding butyrate medium with a cartridge-type peristaltic pump (Master Flex® L/S digital drive, Model 7523-80, Cole-Parmer, Canada) to maintain a hydraulic residence time (HRT) of 2.6 h. After the steady-state current density reached at ~1.15 A/m2 in MxC-3, the electrical conductivity of the biofilm anode was measured. MxC-4 was operated using H2 as the electron donor to confirm if exoelectrogens prefer to consume H2 for current generation; we summarized inoculation and operation of MxC-4 in the Supplementary Material. All experiments were conducted in a temperature controlled room at 25 ± 1°C. 2.3. Serum bottle tests To assess butyrate fermentation without anode respiration of exoelectrogens, biofilms and supernatant 5 mL collected from MxC-1 were transferred to serum bottles having 10 mL of the butyrate medium; the total volume of serum bottle was 20 mL (working volume
15 mL and headspace 5 mL). After sparging the bottles with ultra-pure nitrogen (99.999%) for 1 min, the liquids in the serum bottles were continuously mixed at 100 rpm using a magnetic stirrer (Model 650, VWR International Inc., Canada) in the temperaturecontrolled room. We measured initial and final concentrations of volatile fatty acids in liquid samples. The volume and composition of biogas produced from serum bottles were intermittently monitored. The total duration of the serum bottle test was 72 hours. Serum bottle tests were conducted in duplicates, and average data was used for calculating cumulative H2 production, according to the literature [13].
2.4 Analytical Methods We quantified volatile fatty acids using a gas chromatography (GC) (Model: Hewlett Packard HP 5890 Series II) equipped with a Nukol fused-silica capillary column and flame ionization detector (FID) using He as a carrier gas. The initial temperature of the column was 110°C, increasing to 195°C at the rate of 8°C/min, and then held constant at the final temperature of 195°C for 9 min. Injector and detector temperatures were maintained at 220°C and 280°C, respectively. The liquid samples were acidified to pH ~ 2 using 1 N phosphoric acid prior to GC-FID analyses, and then filtered using 0.2 µm membrane syringe filter. As confirmed in our previous study [9], the detection limits for acetate, n-butyrate, and i-butyrate were 0.03, 0.02, 0.004 mM, respectively.
All liquid
samples were analyzed in triplicates. The volume of biogas produced in serum bottle tests was measured with a friction-free glass syringe (Perfektum Glass Luer Lock Syringe 30 mL, Thomas Scientific, NJ, USA). H2 percentage in the biogas was quantified with a
GC (Model 310, SRI Instruments, Torrance, CA) equipped with a thermal conductivity detector (TCD) and a molecular sieve column (Molesieve 5A, mesh 80/100, 6 ft × 1/8 in.). Argon (99.999%, PraxAir, Canada) was used as a carrier gas with a flow rate of 10 mL/min under a pressure of 21 psi for the GC. The temperatures of the column oven and the detector of the GC were kept at 31°C and 200°C, respectively. The dissolved H2 concentration in MxC-1 and MxC-2 was measured using a H2 microelectrode (Unisense H2-100, Unisense A/S, Denmark) with a tip diameter of 100 m connected with a 4-channel microsensor multimeter (Microsensor Multimeter for Unisense Sensors, 2x pA, 1x mV and 1x Temp channel, Unisense A/S, Denmark). Using 20 mL vials (10 mL deionized water and 10 mL headspace) we prepared different H2 concentrations in liquid and H2 gas compositions in headspace by sparging the vials with H2 and N2 gases (99.999%, PraxAir, Canada) in given times. The vials were vigorously shaken with a vortex mixer for 3 minutes at 3,000 µ at room temperature. The percentage of H2 gas in the headspace was measured with the GC-TCD and converted to the concentration of dissolved H2 in liquid using Henry’s law constant for H2 (0.00078 mol/L-atm at 25°C), giving a range of 13-399 µM of aqueous H2. We then built a calibration curve between the prepared H2 concentrations and the voltage values read by the H2 microelectrode (R2=0.99). We measured the dissolved H2 concentration in the anolyte of MxC-1 by inserting the H2 microsensor through a gas-tight rubber on top of MxC-1; the microsensor was 7 cm distant from the anode in MxC-1. H2 molecules can be consumed by exoelectrogens, methanogens or acetogens in biofilm anodes or bulk liquid, which can bias aqueous H2 measurements with the microsensor. To overcome this challenge, we used MxC-2 and positioned the H2 microelectrode tip in the vicinity of the anode using
a motorized micromanipulator (Unisense MM33, Unisense A/S, Denmark). We then measured dissolved H2 concentration over time using SensorTrace Suite v2.1.100 software in a computer connected to the microsensor multimeter. The electrical conductivity of the anode biofilms was measured using MxC-3 with the two-probe method according to the literature [18–20]. Briefly describing, the gold anodes and the cathode were disconnected temporarily (open circuit mode). Then, a linear sweeping voltage of 0-0.05 V in a step of 0.025 V was applied across two gold electrodes using a source measurement unit (Keithley 2400, Keithley Instruments, Inc., USA), and the current was recorded for each voltage. The voltage ramp was applied in 4-5 cycles until a steady-state current-voltage (I-V) response was obtained. Observed biofilm conductance (GBiofilm (obs), mS) was calculated from steady-state I-V curves. The ionic conductance (Gcontrol, mS) was measured with an abiotic MxC-3 to account for ionic current across the non-conductive gap using the butyrate medium. Intrinsic biofilm conductance was determined by considering ionioc conductance (GBiofilm = GBiofilm (obs) - Gcontrol, mS) and we calculated biofilm conductivity with Eq. (1) [21].
Where, Gbiofilm is intrinsic biofilm conductance (mS) (GBiofilm = GBiofilm (obs)-Gcontrol), Lf is the biofilm thickness (μm), L is the length of the electrodes (1.5 cm), and a is half of the non-conductive gap between two electrodes (25 μm). Biofilm thickness in MxC-3 was measured with a microelectrode as previously described in the literature [14,19]. Supplementary Material provided details of biofilm thickness measurement.
2.5 Microbial Community Analysis We collected biofilm and suspended cells in MxC-1 to analyze microbial community. No methane was detected in this study, and hence we focused on bacterial community analysis. At the end of MxC-1 operation (days 120), we disintegrated MxC-1 in the anaerobic chamber and took a portion of biofilms with a sterilized spatula. We transferred the anolyte into multiple microcentrifuge tubes (~1.5 mL in each tube) and centrifuged them at 5,000 rpm for 2 min (Eppendorf 5424, Hamburg, Germany) to collect pellets. The biofilm and planktonic cells were stored at -20 °C before DNA extraction and sequencing steps. DNA was extracted using the MoBio PowerSoil DNA extraction kit (Solana Beach, CA, USA) according to the manufacturer’s provided protocol. Concentration and purity of DNA were measured using the NanoDrop ND-1000 (NanoDrop Technologies, Wilmington, DE, USA). Typically the values were around 50 ng/μL and around 1.9 (Abs 260 nm/Abs280 nm), respectively. Barcode 454 pyrosequencing targeting bacterial 16S rRNA genes was conducted using the extracted community DNA at the Macrogen (Seoul, South Korea). For the amplification of bacterial 16S rRNA genes, bacterial universal primers (27F: 5´-AGAGTTTGATCMTGGCTCAG-3´, 518R: (5´ATTACCGCGGCTGCTGG-3´) were used [22,23]. The detailed procedure of pyrosequencing was described in a previous study [24]. From sequencing results, species richness (Chao1 and ACE indices) and diversity (Shannon and Simpson indices) were calculated using the Mothur utility [24]. Operational taxonomic units (OTUs) were defined as groups of sequences showing > 97% sequence identities using the Mothur utility. Taxonomic identities of the OTUs were determined based on the SILVA database (https://www.arb-silva.de).
3. Results and Discussions 3.1 Butyrate consumption in a biofilm anode Fig. 2-(a) shows the evolution of current density with time in two consecutive batch operations of MxC-1 after we confirmed four replicate patterns of current density to time. For both runs, the current density rapidly peaked at 7.8-8.8 A/m2 after addition of fresh butyrate medium, and then decreased in a week approximately. Like current density, the concentration of n-butyrate gradually reduced with time, as shown in Fig. 2-(b). i-butyrate (0.17-4.9 mM) and acetate (0.35-14.7 mM) were intermediately accumulated and degraded with time. Isomerization of n-butyrate was commonly found as an intermediate in n-butyrate degradation of methanogenic enrichments [25,26]. The intermittent accumulation of acetate and i-butyrate from n-butyrate suggests that butyrate fermentation to acetate and H2 (oxidative acetogenesis) would occur in MxC-1 (C4H7O2- + 2H2O → 2CH3COO- + H+ + 2H2). This fermentation reaction, however, is thermodynamically unfavorable under standard conditions at pH 7; the standard Gibbs free energy at pH 7 (ΔG') for the n-butyrate fermentation is +48.3 kJ/mol butyrate [1,27]. Aqueous H2 concentration (or partial pressure of H2 (PH2)) should be maintained very low for the butyrate fermentation to occur. Hence, the degradation of n-butyrate and intermittent accumulation of ibutyrate and acetate imply that H2-oxidizing microorganisms would consume H2 produced from the butyrate fermentation, overcoming the thermodynamic bottleneck in the oxidative acetogenesis of butyrate. This interpretation well accords to the literatures [1,9,28], suggesting syntrophic interactions among fermenters, H2 consumers and exoelectrogens in biofilm anodes fed with fermentable substrates.
3.2 Thermodynamic evaluation of butyrate fermentation in the biofilm anode: threshold H2 concentration To drive the butyrate fermentation in MxC-1, dissolved H2 concentration should be less than threshold H2 concentration that is able to trigger butyrate fermentation to acetate and H2 thermodynamically. According to Eq. (2) we computed the threshold PH2 and converted it to the threshold H2 concentration using Henry’s law constant for H2 (0.00078 mol/L-atm at 25°C).
Where, ΔG' is the standard Gibbs free energy for butyrate fermentation (+48.3 kJ/mol butyrate), R is the ideal gas constant (0.00831 kJ/mol-K), T is the temperature (K), [HBu] is the total butyrate concentration (M) including n-butyrate and i-butyrate, [HAc] is the acetate concentration (M), and [H+] is the proton concentration (M). For thermodynamic calculation, we assumed that i-butyrate is transformed to n-butyrate and the n-butyrate is fermented to acetate and H2 [26,29]; energy change in isomerization of n-butyrate is negligible [25], so we ignored small energy change in butyrate isomerization for this computation. Fig. 3 shows the evolution of the threshold H2 concentration in MxC-1 when n-butyrate, i-butyrate, and acetate concentration were varied with time (Fig. 2). Thermodynamic evaluation on the butyrate fermentation indicates that the threshold H2 concentration should range from 0.1 to 7.2 μM (PH2 10-942 Pa), indicating that H2 gas should not be accumulated in the anodic headspace of MxC-1. We did not detect any H2 gas in the headspace with the GC-TCD (detection limit > 0.01 atm H2), and this result accords to the threshold H2 calculation in Fig. 3. We measured the dissolved H2 concentration in the anolyte of MxC-1 with the H2 microsensor that
can precisely quantify a few μM ranges of dissolved H2 with a detection limit of 0.3 µM (www.unisense.com). Unexpectedly, no dissolved H2 was detected in the anolyte, which contradicts to the thermodynamic analysis of threshold H2 computation. The microsensor was located at 7 cm distant from the anode in MxC-1. H2 molecules produced from the butyrate fermentation could be scavenged by H2 consumers in the biofilm anode or bulk liquid before the H2 reaches the microsensor; in Section 3.4 we further explored dissolved H2 concentration in the butyrate fermentation using MxC-2 where the microsensor physically touched the anode surface to mitigate H2 consumption effect during measurements. Hydrogenotrophic methanogens, H2-oxidizing exoelectrogens, or acetogens can consume H2 produced from butyrate fermentation. We did not detect any methane gas in the headspace of MxC-1 during experiments, excluding the methanogens from H2 consumer candidates. To test H2 oxidation by exoelectrogens, we operated MxC-4 using H2 gas as the sole electron donor, but current density in the H2-fed MxC was negligible at 0.03-0.24 A/m2 close to endogenous decay current density (see Supplementary Material for details). Thus, direct H2 oxidation by exoelectrogens would be negligible for H2 consumption in MxC-1. It is likely that acetogens would be main H2 consumers in MxC-1, suggesting significance of the syntrophy between acetogens and exoelectrogens for current generation (H2 + CO2 → acetate → electrical current). Several studies proposed this syntrophic interaction in MxCs fed with fermentable substrate [1,9,12]. In serum bottle test, the concentration of n-butyrate slightly decreased in 72 h, along with small increase of i-butyrate and acetate, as described in Table 1. Cumulative H2 production was small at 0.14±0.03 mL H2 and its production was stopped at 48 h. No methane was detected during the experiments, consistent to MxC-1, and the concentration of the acids was not changed after 72 h of reaction
time (data not shown). The evolution of VFAs and H2 in serum bottle tests supports oxidative acetogenesis of n-butyrate (butyrate conversion to acetate and H2) in MxC-1, but the cease of H2 production and n-butyrate degradation proves significance of exoelectrogens and their syntrophy with fermenters and H2 consumers for continuing the oxidative acetogenesis.
3.3 Microbial community in biofilm and planktonic cells From the two samples (biofilm and suspended cells in MxC-1) 1,036 – 1,450 of sequences were used for bacterial community analysis after removal of chimera and short sequences (< 250 base pairs). Table S1 compares species (OTU) richness and diversity estimated from the retrieved 16S rRNA gene sequences. Both two species richness indices (Chao1 and ACE) estimated lower richness of bacteria species in the biofilm than planktonic cells. The same pattern was also observed in two species diversity indices (Shannon and Simpson). Bacterial community compositions were compared in Fig. 4. Communities were not significantly different at the phylum level (see Fig. 4(a)). The most dominant phylum was Proteobacteria for the biofilm and planktonic cells (61.3 – 90.5%), following Firmicutes, Synergistetes, Bacteroidetes, and Spirochaetes. However, communities were very different at the genus level, as shown in Fig. 4(b). Geobacter was predominant (83.6% out of total sequences) in the biofilm anode, like the mother MxC. Small populations of Sphaerochaeta and Treponema, that can build the syntrophy with butyrate fermenters as acetogens [30,31], were identified along
with 11.85% unclassified bacteria. This result indicates that Geobacter is the main exoelectrogen in the biofilm anode, as reported by the literature [9,18–20,32]. The biofilm community supports the proposed syntrophy between exoelectrogens and acetogens. In comparison, Geobacter only comprised 2.1% and the other genera were dominant in planktonic cells: Pseudomonas (18.7%), Aeromonas (16.4%), Sphaerochaeta (8.2%), Treponema (3.8%), and Proteiniphilum (3.5%). Metabolic functions of Pseudomonas and Aeromonas are versatile [33,34], and hence it is challenging to assess their metabolic functions with 16S rRNA genes. Pseudomonas and Aeromonas can work as fermenters [35,36] or conduct anaerobic respiration using conductive solids [37,38]. For instance, some Pseudomonas species can produce electron shuttling compounds (e.g. pyocyanin) for mediated electron transport to solid conductors [38,39]. Both Pseudomonas and Aeromonas species also possess genes for EET and can produce the electrical current in MxCs [37,40,41]. Consistent to the biofilm, Sphaerochaeta and Treponema were also identified in the suspended cells, acetogen candidates. The bacterial community of the planktonic cells implies that different exoelectrogens might play a role of acetate consumption in bulk liquid, but consistently supports the proposed syntrophy of acetogens and exoelectrogens. 3.4 H2 production and consumption in the butyrate-fed biofilm anode Fig. 5 shows the change of current density and dissolved H2 concentration with time in MxC-2. Like the batch run of MxC-1, current density gradually increased, reached a plateau, and decreased with time. Dissolved H2 concentration was as high as 12.4 µM (PH2 1,640 Pa) at the initial fermentation of n-butyrate, which indicates H2 production in oxidative acetogenesis of butyrate. Dissolved H2 concentration decreased with time, and it became as small as 3.5 µM at the end of operation (140 h). This result clearly evidences H2 consumption in MxC-2; here, we only sampled and analyzed liquid samples at the end of batch run because sampling readily disturbed
the measurements of dissolved H2 concentration with the microsensor. The measured H2 concentration was very close to 3.1 µM of threshold H2 concentration calculated with Eq. 2 (n-butyrate 2.6 mM, i-butyrate 1.3 mM, and acetate 0.9 mM), validating the thermodynamic evaluation of butyrate fermentation and threshold H2 calculation discussed early in Section 3.2. Abrupt increase of current density involved sharp decrease of dissolved H2 concentration in the biofilm anode (an area highlighted with blue color in Fig. 5). This experimental result first proves the significance of H2 consumption for exoelectrogens to generate electric current in the butyrate-fed biofilm anode. Based on molecular biology data, the syntrophic interaction between exoelectrogens and acetogens is a key to overcome a thermodynamic barrier in utilization of butyrate in the biofilm anode. 3.5 Biofilm conductivity Fig. 6 shows the biofilm conductivity measured with MxC-3. The biofilm conductivity was relatively stable at 0.67±0.14 mS/cm in 12 measurements for consecutive four days (biofilm thickness 138±19 μm). The relatively large standard deviation in the average biofilm conductivity would indicate the requirement of a more robust method on biofilm conductivity. However, this biofilm conductivity is within the range of biofilm conductivities (0.25-5 mS/cm) observed for Geobacter-enriched biofilm anodes [14,16,17,19]. This experimental result first proves that the butyrate-fed biofilm is electrically conductive, although complex syntrophic interactions among fermenters, H2 consumers and exoelectrogens can be built in the biofilm.
This high biofilm
conductivity means that electrical conduction governs EET to the anode in the biofilm. According to Eq. (3), the maximum current density achievable in the EET step is computed at ~ 30 A/m2. This calculated current density is even several times higher than the highest current density in acetate-fed biofilm anodes [18,42,43].
j = Kbio (Eanode – EEC° )/Lf
(3)
Where, Kbio is biofilm conductivity (S/m), Eanode is anode potential (V) (-0.2 V vs. SHE), and EEC° is the standard potential for a ratelimiting extracellular cofactor at pH 7 (V). We assumed EEC° = -0.26 V vs. SHE for calculating the maximum current density [18,44]. Low current density (< 2 A/m2) has been commonly observed in MxCs using butyrate as substrate [45,46], but we have not well understood which step causes such low current density in butyrate-fed MxCs. Our study indicates that intra and intercellular electron transfer would mainly limit current density in butyrate-fed biofilms, not EET. The kinetic assessment of intracellular and intercellular electron transfer in the butyrate-fed biofilm is not straightforward, due to complex microbial syntrophy including butyrate fermentation, acetate oxidation, H2 consumption, and mass transfer of intermediate metabolites (e.g., H2 and acetate transfer). More study is required to investigate a rate-limiting step in the intracellular and intercellular electron transfer kinetics for improving current density in butyrate-fed MxCs. The electrically conductive biofilm implies that direct interspecies electron transfer (DIET) might occur among butyrate fermenters, H2 consumers, and exoelectrogens, as proposed by recent studies [47,48]. DIET is an efficient manner to build electronsharing networks among microorganisms [48–51], but our study proved the significance of H2 syntrophy in the electrically conductive biofilm anode. For inter-electron transport, DIET may be better than hydrogen molecules considering mass transport limitation of H2.
In energetic point of view, however, H2 syntrophy would be superior to DIET because H2 is a great energy-carrier to syntrophic partners; in comparison, no studies have proved energy benefit to microorganisms in DIET.
4. Conclusions A high electrical conductivity (0.67±0.14 mS/cm) was measured for the butyrate-fed biofilm, which demonstrates that EET does not limit current density in the biofilm. Despite of high electrical conductivity in the biofilm, in-situ measurements of H2 concentration confirmed H2 production and consumption consistent to current generation in the butyrate-fed biofilm. Geobacter was identified as the main exoelectrogen in syntrophy with Sphaerochaeta and Treponema, supporting the significant roles of acetogens as the H2 scavenger and syntrophic partner in the biofilm. The high electrical conductivity of the biofilm suggests that DIET might occur to link exoelectrogens with syntrophic partners for butyrate utilization, but this study proves the significance of H2-based syntrophy even in the electrically conductive biofilm. Acknowledgements This research was funded by Natural Sciences and Engineering Research Council of Canada Discovery Grant (RGPIN-2016-04155). We appreciate Dr. Jangho Lee’s supports to calibration of the H2 microsensor. References
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Table and figure captions Table 1. Changes of acids and biogases in serum bottles lacking anode respiration.
Fig. 1. Schematic diagram showing configuration of (a) MxC-1 and MxC-4, (b) MxC-2, and (c) MxC-3. Note. MxC-2 and MxC-3 were designed for quantifying dissolved hydrogen concentration and biofilm conductivity during butyrate fermentation, respectively. Figures are not drawn to scale. Fig. 2. Profiles of current density and main VFAs to time in MxC-1. (a) Current density, and (b) main VFAs of n-butyrate, i-butyrate, and acetate.
Fig. 3. The threshold H2 concentration for butyrate fermentation to acetate and H2 in MxC-1. The H2 concentration was computed with Eq. (2). Fig. 4. Relative abundance at (a) phylum, and (b) genus level. Fig. 5. Current density and dissolved hydrogen concentrations on anode surface in MxC-2 during fed-batch operation. Blue highlighted areas indicate abrupt increase of current density and decrease of dissolved H2 concentration in MxC-2. Fig. 6. Current density and measured biofilm conductivity during operation of MxC-3.
Table 1. Changes of acids and biogases in serum bottles lacking anode respiration.
Initial
Final
n-butyrate
10.8±0.1 mM
8.2±1.8 mM
i-butyrate
0.21±0.1 mM
0.8±0.2 mM
Acetate
0.6±0.3 mM
1.7±1.0 mM
H2 gas
0
0.14±0.03 mL
CH4 gas
0
N.D.
Serum bottle tests were conducted in 72 h. N.D.: not detected.
Highlights
The electrical conductivity of the butyrate-fed biofilm was as high as 0.67 mS/cm H2-based syntrophy played an important role in the electrically conductive biofilm Aqueous H2 concentration decreased as current increased Acetogens (Sphaerochaeta and Treponema) were primary H2 consumers in the biofilm Intracellular electron transport mainly limited current in the biofilm
Fig. 1. Schematic diagram showing configuration of (a) MxC-1 and MxC-4, (b) MxC-2, and (c) MxC-3. Note. MxC-2 and MxC-3 were designed for quantifying dissolved hydrogen concentration and biofilm conductivity during butyrate fermentation, respectively. Figures are not drawn to scale.
Fig. 2. Profiles of current density and main VFAs to time in MxC-1. (a) Current density, and (b) main VFAs of n-butyrate, i-butyrate, and acetate.
34
Fig. 3. The threshold H2 concentration for butyrate fermentation to acetate and H2 in MxC-1. The H2 concentration was computed with Eq. (2).
35
Fig. 4. Relative abundance at (a) phylum, and (b) genus level.
36
Fig. 5. Current density and dissolved hydrogen concentrations on anode surface in MxC-2 during fed-batch operation. Blue highlighted areas indicate abrupt increase of current density and decrease of dissolved H2 concentration in MxC-2.
37
Fig. 6. Current density and measured biofilm conductivity during operation of MxC-3.
38