Biohydrogen production via biocatalyzed electrolysis in acetate-fed bioelectrochemical cells and microbial community analysis

international journal of hydrogen energy 33 (2008) 5184–5192

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Biohydrogen production via biocatalyzed electrolysis in acetate-fed bioelectrochemical cells and microbial community analysis Kyu-Jung Chae, Mi-Jin Choi, Jinwook Lee, F.F. Ajayi, In S. Kim* Department of Environmental Science and Engineering, Gwangju Institute of Science and Technology (GIST), 261 Cheomdan-gwagiro, Buk-gu, Gwangju 500-712, South Korea

article info

abstract

Article history:

Hydrogen was efficiently produced via acetate oxidation using a two-chambered bioelec-

Received 10 May 2008

trochemical cell (BEC), a modified microbial fuel cell (MFC), in which a cathode was kept

Accepted 26 May 2008

free of oxygen and the thermodynamic barrier overcome by augmenting the electrochem-

Available online 3 August 2008

ical potential achieved by bacteria with the addition of a small external voltage. The production of hydrogen gradually increased with increasing applied voltage from 0.1 to

Keywords:

1.0 V, reaching its maximum efficiency of 52.5% at 0.8 V, corresponding to a hydrogen yield

Bacterial community

of 2.1 mol per 1 mol acetate. The anodic loss of electrons was more detrimental compared

Biocatalyzed electrolysis

to that of cathodic loss, especially with the higher applied voltage range. The cathode head-

Hydrogen

space gas consisted mainly of hydrogen (>96.6%), but with minor methane (2.5%) and car-

Microbial fuel cells

bon dioxide (0.9%) impurities diffused from the anode chamber. To maintain the purity or prevent the loss of the hydrogen produced, much concern is required for the control of CO2 and CH4 because these gases diffuse more readily through a Nafion 117 membrane than H2. To produce hydrogen, the continuous augmentation of the circuit by an external voltage had no harmful effect on the bacterial viability but resulted in a remarkable change in the bacterial community. There was a substantial decrease in species diversity with a single emergent Pelobacter propionicus-like species in the BEC. Interestingly, Geobacter-like species were integral members of the bacterial consortia in an MFC and BEC. ª 2008 International Association for Hydrogen Energy. Published by Elsevier Ltd. All rights reserved.

1.

Introduction

The production of hydrogen via current anaerobic dark fermentation is limited to a maximum of 4 mol of hydrogen per mole of glucose, with the fermentation end product (acetate; 2 mol/mol glucose) that is not further converted to hydrogen by bacteria due to the endothermic nature [1,2]. However, this thermodynamic barrier can be overcome by generating hydrogen from acetate using a modified microbial fuel cell

(MFC). The MFC generates electricity, but can be modified to produce hydrogen instead, as shown in two half reactions indicated below (Eqs. (1) and (2)). This modified MFC, recently suggested and referred to as biocatalyzed electrolysis [3,4] or the bio-electrochemically assisted microbial reactor (BEAMR) process [1] or electrohydrogenesis [5], has been considered as a promising new technology for the production of biohydrogen from organics. However, in order to produce hydrogen using this modified MFC, the cathode is kept free of oxygen

* Corresponding author. Tel.: þ82 62 970 2436; fax: þ82 62 970 2434. E-mail address: [email protected] (I.S. Kim). 0360-3199/$ – see front matter ª 2008 International Association for Hydrogen Energy. Published by Elsevier Ltd. All rights reserved. doi:10.1016/j.ijhydene.2008.05.013

international journal of hydrogen energy 33 (2008) 5184–5192

and the thermodynamic barrier must be overcome, as the direct production of hydrogen and carbon dioxide from the hydrolysis of acetate is not thermodynamically feasible. Anode : CH3 COOH þ 2H2 O/2CO2 þ 8Hþ þ 8e : E0  0:28 VðNHEÞ Cathode : 8Hþ þ 8e /4H2

(1) : E0  0:42 VðNHEÞ

(2)

At pH 7, the equilibrium potentials for the oxidation of acetate (1 M) at the anode and the reduction of protons to hydrogen at the cathode are 0.28 and 0.42 V (NHE), respectively. Therefore, hydrogen can theoretically be produced at the cathode by applying a circuit voltage greater than 0.14 V (i.e., 0.42 to 0.28 V), as previously described by Rozendal et al. [3]. The required voltage (0.14 V) is substantially lower than that needed for the production of hydrogen from the direct electrolysis of water, which is theoretically 1.23 V at neutral pH, but practically requires above 1.8 V under alkaline conditions due to the diverse overpotentials. The modified MFC has functional advantages over the technologies currently used for generating hydrogen from organic matter; firstly, the direct conversion of substrate energy to hydrogen enables high conversion efficiency. In addition, the high purity (100% in theory) of the hydrogen produced is another advantage compared to other biohydrogen production methods from organic materials containing a diverse mixture of biogases (e.g., CH4, CO2, H2S, NH3, etc.) and other impurities and, therefore, require expensive gas purification [6,7]. Liu et al. [1] achieved a yield of 2.9 mol H2/mol acetate (¼72.5% hydrogen recovery) in their feasibility study using BEAMR process by supplying 0.85 V. Rozendal et al. [3] obtained an overall hydrogen production efficiency of 53% in their acetate-fed biocatalyzed electrolysis system at an applied voltage of 0.5 V. Furthermore, based on the optimization of the rector configuration to reduce the internal resistance, Rozendal et al. [4] suggested a single chamber biocatalyzed electrolysis by implementing a gas diffusion electrode in two configurations: with both cation and anion exchange membranes (CEM and AEM). The performances of both configurations were comparable with overall hydrogen production efficiencies around 23% at 1.0 V. Unlike single chamber MFCs which show low internal resistance by removing the CEM [8], in

A 2

3

5

5185

biocatalyzed electrolysis the presence of a membrane is essential to separate the hydrogen produced at the cathode from the biogas mixture in the anode chamber. Biocatalyzed electrolysis also applies electrochemically active bacteria which are capable of using an electrode as an external electron acceptor for the oxidation of organic compounds [9,10]. Therefore, electrons and protons are retrieved at the anode in the same manner as with MFC, except that an external voltage is supplied continuously to reduce the protons to hydrogen at the cathode. This exogenous voltage augmentation can affect the consortia or viability of bacteria, but there has been no study reporting this matter. Furthermore, much of the information required is still unclear, such as individual electrode response to the voltage applied, the governing overpotentials associated with the electrode reactions, and the gas permeability through the CEM, etc. In our study, this modified MFC was termed a bioelectrochemical cell (BEC), even though others have referred to this as biocatalyzed electrolysis or BEAMR process. In this study, the performances of hydrogen evolution using an acetateenriched BEC, and the bacterial diversity using 16S rDNA sequencing, as well as their viability on the anode, were collectively investigated. The gas permeability through the CEM was also examined to determine the loss of hydrogen from the cathode to the anode compartment, as well as the contamination by the diffusion of gas from the anode and vice versa.

2.

Materials and methods

2.1.

Bioelectrochemical cell (BEC) design and operation

Two-chambered BECs (180 mL working volume each) were constructed, as previously described [11], with a Nafion 117 membrane (a projected area of 25 cm2, DuPont Co., USA) and 3 mm electrode spacing (Fig. 1). The anode consisted of a carbon felt electrode (25 cm2, 6 mm thickness, Morgan, UK), but the cathode was made of a perforated titanium plate (Labco Co., South Korea), the same size as the anode, with a 0.5 mg/ cm2 platinum coating as a catalyst. Both the anode and cathode chambers were equipped with an Ag/AgCl reference

B

1

4

Fig. 1 – The two-chambered BEC used in this study. (A) Schematic diagram: (1) anode, (2) Nafion 117 membrane, (3) cathode, (4) edge-sealing gasket, (5) reference electrode. (B) Photograph.

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electrode (þ0.2 V vs. NHE; Microelectrode, USA) to measure the separate electrode potentials. As an external power source a potentiostat (BAS Epsilon, Bioanalytical systems, Inc., USA) was used. A voltage in the range of 0.1–1.0 V was applied to the circuit by connecting the working electrode of the potentiostat to the anode, with both the counter and reference electrode connected to the cathode. The anode and cathode potentials were measured using a multimeter (Keithley, OH, USA) and reference electrode (Ag/AgCl). All reported potentials were reported against the Ag/AgCl reference, except where otherwise noted. The anode chamber was filled with an autoclaved anaerobic nutrient mineral buffer (NMB, pH 7.0) solution and anaerobic digester sludge (20% v/v), whereas, the cathode chamber contained a phosphate buffer (50 mM, pH 7.0), as described previously [11]. Acetate (2.0 mM) was used as the sole substrate for the entire run. Both chambers were initially purged with nitrogen gas to remove oxygen, and mixed gently with a magnetic stirrer.

2.2. Enrichment of electrochemically active microorganisms For the enrichment of electrochemically active microorganisms, BECs were operated in the conventional MFC mode with oxygen as an electron acceptor in the cathode for over 5 months, and then changed to the hydrogen production mode, as described above. The BECs were run in a temperature controlled room at 26–28  C, in a fed-batch mode, with the feed solution in the anode chamber replaced as the anode potential decreased (became less negative) to a value of 50 mV (vs. Ag/AgCl). All performance evaluations were conducted once the system had become fully acclimated to the substrates and demonstrated repeatable cycles of hydrogen generation.

2.3.

Calculations and analyses

The individual anode and cathode potentials, as well as the overall cell potential (anode–cathode), were continuously monitored at 10 min intervals using a multimeter connected to a personal computer. The Coulombic efficiency (CE) was calculated, as previously described [12]. For the CE and overall hydrogen recovery analyses, the background current (current at the working electrode in the absence of substrate) was determined for each experiment and then subtracted from all values prior to calculating the total electron recovery. The acetate concentration in the solution was regularly measured for calculation of the CE using a gas chromatograph (Hewlett Packard 6890 plus series) equipped with a flame ionization detector (FID) and an EC-1000 capillary column (Alltech, USA) with helium as the carrier gas. The gas compositions from the anode and cathode were regularly analyzed using a gas chromatograph (Hewlett Packard 5890 plus series) equipped with a thermal conductivity detector (TCD) and a molecular sieve column (Alltech Molesieve 5A 80/100) with nitrogen as the carrier gas.

2.4.

Bacterial community analyses

The enriched biofilm was detached from the anode surface in the two-chambered BECs fed with acetate using a sterilized

razor knife. Total genomic DNA was extracted from the collected biofilms using a DNA extraction kit (AccuPrep Genomic DNA extraction kit k-3032, Bioneer, Republic of Korea), in accordance with manufacturer’s instructions, and used as template DNA in the PCR amplification. Two universal bacteria primers [13], 9f (50 -GAG TTT GAT CCT GGC TCA G-30 ) and 1512r (50 -ACG GTA CCT TGT TAC GAC TT-30 ), were used for 16S rRNA gene fragment (1504 bp) amplification. PCR amplification was performed in a mastercycler (Eppendorf, Germany), with a thermal cycle program, including an initial enzyme activation step of 6 min at 95  C, followed by 30 cycles. Each cycle consisted of denaturation of 60 s at 94  C, annealing of 60 s at 58  C and extension of 90 s at 72  C. The 16S rRNA gene from the PCR product was purified using an AccuPrep PCR purification kit (Bioneer, Republic of Korea) to construct gene cloning using a yT&A cloning vector kit (RBC, Taiwan). Ligation of the purified 16S rRNA to the yT&A cloning vector (25 ng/mL) was carried out overnight, in the dark, at 4  C using YEA T4 DNA ligase (3 U/mL). For transformation, 5 mL aliquots of ligation reaction mixture were mixed with 50 mL of competent cells (High 108 HIT-DH5a, RBC, Taiwan). After placing the transformation reaction mixture in a 42  C water bath for 45 s to induce heat shock, it was transferred on an LB ampicillin (70 mg/L) agar spread with X-gal/IPTG (40 mg/mL). The transformed cells were incubated overnight at 37  C for blue/white screening, with white colonies chosen and suspended in 100 mL PBS buffer for purification of plasmid DNA. PCR amplification was performed on each purified plasmid using M13 primers, forward primer 50 -GTT TTC CCA GTC ACG AC-30 and reverse primer 50 -TCA CAC AGG AAA CAG CTA TGA C-30 . Each plasmid DNA from the PCR product was purified again, and then sequenced automatically with an ABI Prism BigDye Terminator cycle sequencing ready reaction kit (Applied Biosystems, USA) and DNA sequencer (model 3700, Applied Biosystems, USA). The primer, 805R (50 -GAC TAC CAG GGT ATC TAA T-30 ), was used for half sequencing. The sequenced 16S rRNA gene was analyzed using the NCBI search tool to identify the closest matching sequence. Sequences were compared to those in the GenBank databases using the Basic Local Alignment Search Tool (BLAST) algorithm to determine the approximate phylogenetic affiliations.

2.5.

Bacterial viability assay

In order to evaluate the cell viability of the biofilm on the anode in the BECs, a LIVE/DEAD BacLight Bacterial Viability Kit (Molecular Probes, Inc., Eugene, OR, USA), which employs two nucleic acid stains: the green-fluorescent SYTO 9 stain for live cells and the red-fluorescent propidium iodide stain for dead cells, was used to assess the cell membrane integrity. All steps, including sample preparation and staining, were conducted under anaerobic conditions to avoid oxygen damage prior to the observation using a confocal laser scanning microscope (CLSM, LSM 5 PASCAL, Carl ZEISS, Inc., USA). Live (in green fluorescence) and dead bacteria (in red) were observed separately using fluorescence microscopy, and then the monochrome images of a series of the pin floc or dispersed live and dead cells were quantified using the image analysis program of the CLSM.

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3.1. Current densities and electrode potentials as a function of applied voltage

3.5

Current (mA)

Current density (A/m2)

3.0

Without HAc HAc (2.0 mM)

2.5 2.0 1.5 1.0 0.5 0.0 0.0

0.2

20

0.4

0.6

0.8

1.0

Applied voltage (V) Fig. 2 – Current densities as functions of the applied voltage in a two-chambered hydrogen generating BEC run for over 5 months. The current generated and the electrode potentials were measured in both the absence and presence of substrate (acetate 2.0 mM). Data represent the means ± SD in two separate batch experiments in duplicate.

2.5

A

H2 HAc

15

2.0 1.5

10 1.0 5

0.5

0 2.5

Acetate (mM)

To evaluate the cell performance of the BEC, and confirm the electron source, the current densities as functions of the applied voltage were measured using two-chambered hydrogen generating BECs which had been run for over 5 months for the production of hydrogen after enrichment of the electrochemically active bacteria in the previous 5 months as a normal MFC. Fig. 2 shows the applied voltage scans obtained in duplicate by means of chronoamperometry by stepwise increasing the applied voltage from 0.0 to 0.8 V using a potentiostat. The current generated was automatically logged every 10 s for 1 h for each applied voltage step, and the average value then plotted. In the presence of acetate (2.0 mM), the current density significantly increased with increasing applied voltage from 0.04 A/m2 anode surface (at 0.1 V) to 2.91 A/m2 (at 1.0 V) (Fig. 2). To confirm the current generated in the applied voltage scans had indeed originated from the oxidation of acetate, the system was operated in the same manner, but without acetate in the anode medium. In this case, current generated was negligibly low (0.38–0.70 A/m2), with the production of no hydrogen at the cathode. This result indicated the measured current definitely arose from the oxidation of acetate in the anode chamber. In addition, there was no hydrogen production from the BEC with a virgin anode (data not shown). Overall, the electrons are retrieved from acetate oxidation as a result of anodic biocatalysis of the attached electrochemically active bacteria. In the absence of acetate, electrons suddenly charged the surface of the electrodes as a potential was applied, without passing through the external circuit due to the consumption of no electrons at the cathode

as no protons were delivered via the Nafion, consequently the cathodic reaction (8Hþ þ 8e / 4H2) could not be completed. Therefore, the current densities were significantly lower in the absence of acetate, and even at the higher applied voltage range did not increase compared to those obtained in the presence of acetate. Interestingly, the anode potential was more susceptible to increases in the applied voltage, i.e., a substantial decrease (negative value becomes less negative) at the anode, but remained relatively constant at the cathode (data not shown). Therefore, especially at higher applied voltages, the anode potential became less negative than that theoretically required for acetate oxidation (0.28 V vs. NHE), consequently results in the reduction of the overall hydrogen evolution efficiency. Fig. 3 represents the hydrogen gas, current generation and electrode potential response profiles when acetate was fed into the anode chamber of the BECs operating at an applied voltage of 0.3 V. Injected acetate was rapidly degraded, with a typical batch cycle requirement of 46 h, for the almost complete consumption (>98%) of 2.0 mM, which resulted in the corresponding hydrogen and current generation (Fig. 3a and b). The anode potential suddenly increased from 0.220 to 0.410 V, but stabilized over 48 h, and then decreased again as a result of the complete consumption of acetate (Fig. 3c). The anode potential exactly reflected the status of substrate oxidation, therefore, used as an indicator to represent the time to feed in batch operation.

Hydrogen (mL)

Results and discussion

Voltage (V) vs. Ag/AgCl

3.

0.0

B

2.0 1.5 1.0 0.5 0.0 0.3

C Overall cell Anode Cathode

0.0 -0.3 -0.6 0

10

20

30

40

50

60

70

Time (hr) Fig. 3 – (A) Consumption of acetate in the anode chamber and the amount of hydrogen accumulated in the cathode chamber. (B) Current between both electrodes. (C) Responses of the anode and cathode potentials on the addition of 2.0 mM acetate into the anode chamber of the BEC, with an applied voltage of 0.3 V.

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3.2. Coulombic efficiency, cathodic hydrogen efficiency and overall hydrogen efficiency Fig. 4 shows the CE (acetate to e in the anode), cathodic hydrogen efficiency (e to H2 in the cathode) and overall hydrogen efficiency (acetate to H2) in the applied voltage ranges from 0.1 to 1.0 V. An applied voltage between 0.5 and 0.8 V was determined as the optimum for the production of hydrogen, where the recovery of electrons from acetate as hydrogen was over 45.2%. Hydrogen gas was generated at the cathode when the applied voltage was greater than 0.30 V. Even though hydrogen can theoretically be produced at the cathode by augmenting a circuit voltage greater than 0.14 V [1,3], greater voltages are needed in a practical situation due to the overpotential at the electrode. Below an applied voltage of 0.2 V, hydrogen was produced only in negligible amounts. The production of hydrogen gradually increased with increasing applied voltage, reaching a maximum efficiency of 52.5% at 0.8 V, corresponding to a hydrogen yield of approximately 2.1 mol per 1 mol acetate, as a maximum possible production of 4 mol H2/M acetate was assumed. This corresponded to a volumetric hydrogen production rate of approximately 0.052 m3 H2/m3 reactor liquid volume/day, which is 2.6 times faster than the 0.02 m3 H2/m3 reported by Rozendal et al. [3]. Interestingly, however, the overall hydrogen efficiency declined but with the maximum cathodic efficiency when the applied voltage was further increased up to 1.0 V. As the applied voltage was increased from 0 to 1.0 V, the cathode maintained a relatively constant potential, while the anode potential sharply decreased from 0.467 V to 0.053 V in order to sustain the potential difference between the anode and cathode as a result of the externally applied voltage. This shift in the anode potential to an unfavorable range allowed the bacteria to use the respiratory chain in an oxidative metabolism or inhibits the electrochemically active bacteria growing at anode potentials under 0.2 to 0.28 V (vs. NHE). This may provide other acetate utilizing microorganisms with

a competitive advantage over electrochemically active bacteria, consequently resulting in a decreased CE, which consecutively lowers the overall hydrogen recovery at this voltage. Conversely, with applied voltages below 0.3 V, a low cathodic efficiency limited the overall hydrogen efficiency. The CEs, the recovery of total electrons in the substrate as current, were quite dependant on the applied voltage, and ranged from 48.0 to 68.4% at 0.3–0.8 V. This CE range is in the middle of that obtained in a similar system. A CE of 60– 78% was achieved by Liu et al. [1] at applied voltages above 0.25 V. Rozendal et al. [3] reported a notable CE of 92  6.3% at an applied voltage of 0.5 V, but the overall hydrogen recovery was 53  3.5% due to the low cathodic hydrogen efficiency of 57  0.1%, but recently reported a significantly low CE of 22.8  0.2% using a similar system [4]. The cathodic hydrogen efficiency was obviously better at the higher applied voltage range, with up to 92% at 1.0 V. The better cathodic efficiency at higher voltages obtained in this study was consistent with the finding that diffusional hydrogen loss from the cathode to the anode chamber can be reduced by increasing the current densities, as suggested by Rozendal et al. [4]. In order to verify the origin of the hydrogen gas produced, all batch experiments were performed with a control reactor containing the same composition, but without acetate as the substrate, to demonstrate the possibility of hydrogen production by other electrochemical reaction, such as water splitting (Fig. 5) [14,15]. Using the same procedure, but without acetate in the anode medium, the production of hydrogen was negligible, indicating the electrons used for the production of hydrogen were from acetate degradation rather than from water splitting. In the BEC systems, the anodic loss (or overpotential) of electrons was more detrimental than the cathodic loss, especially in the higher applied voltage range. Therefore, more attention to the reduction of the anodic overpotential will be required to achieve greater production of hydrogen. With regard to the governing overpotential for the BEC-like systems, controversial findings have been reported: Rozendal et al. [4] and Liu et al. [1] reported the anodic overpotential,

80

20

60

16

0.16 0.12 0.08 0.04 0.00

Hydrogen (mL)

Conversion efficiency (%)

100

40 20 CE Cathodic Efficiency Overall H2 Efficiency

0

12

0.0 0.2 0.4 0.6 0.8 1.0 1.2

8

HAc 2.0 mM Without HAc

4 0

0.0

0.2

0.4

0.6

0.8

1.0

1.2

Applied voltage (V) Fig. 4 – Coulombic efficiency (acetate to eL in the anode), cathodic hydrogen efficiency (eL to H2 in the cathode) and overall hydrogen recovery (acetate to H2) as functions of the applied voltage (0.1–1.0 V) in BECs fed with 2.0 mM acetate. Bars indicate SD.

0.0

0.2

0.4

0.6

0.8

1.0

1.2

Applied voltage (V) Fig. 5 – Comparison of hydrogen production in the presence and absence of acetate. The inset shows the same measurement without acetate on an adapted scale.

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The gas produced in the anode compartment consisted of 73.6% CO2, 26.4% CH4 and 0.0% H2, while the cathode headspace gas consisted of mainly hydrogen (96.6%), but with minor impurities of methane (2.5%) and carbon dioxide (0.9%) which probably diffused from the anode compartment. Even though, in theory, electrochemically active bacteria can completely oxidize organics to carbon dioxide, methane was still produced in large amounts in the anode compartment. This indicates acetate consumption by methanogens, non-electricity-generating bacteria, resulting in a low overall hydrogen efficiency. In conventional MFC using oxygen as an electron acceptor in the cathode, oxygen leakage into the anode chamber can either lower the CE due to the substrate loss from aerobic respiration by facultative bacteria, or inhibit the growth of obligate anaerobes [11,16,17]. However, although this oxygen associated CE loss is avoidable as the cathode chamber is under anaerobic condition, the CEs obtained in the BEC under various operating conditions were 48.0–68.4% (in the applied voltage of 0.3–1.0 V), which were lower than the expected. These lower CEs mainly resulted from substrate loss by methanogens competing with the electrochemically active microorganisms for the substrate, as confirmed by the large presence of methane (26.4%) in the anode chamber. This result indicates that a specific suppression for methanogens is strongly required to enhance the overall hydrogen recovery. In the BEC, the presence of a membrane, such as Nafion, is essential for preventing the hydrogen gas produced at the cathode from mixing with the gas produced in the anode compartment. In previous studies, a large part of the hydrogen produced at the cathode was suggested to be lost via diffusion from the cathode into the anode chamber through the Nafion membrane, which was based on calculations rather than actual measurements [3]. However, undesirable gas (e.g., CH4, CO2) diffusion from the anode to the cathode compartment, reducing the purity of hydrogen, has not been reported. Hence, gas diffusion through Nafion 117 was investigated in this study. For this, after flushing both chambers of a normally working BEC with nitrogen gas, a standard gas mixture (CO2, CH4, and H2) was filled into only the cathode chamber, and then both chamber identically pressurized to 80 mm H2O using nitrogen gas. Both chambers contained 50% (v/v) liquid medium (NMB for the anode and PBS for the cathode chamber) to prevent the Nafion membrane from drying. The gas contents in both chambers were then measured over time at a constant temperature, but without an applied voltage. As shown in Fig. 6, all gases tested including CO2, CH4 and H2 diffused through the Nafion 117 membrane. Of these, carbon dioxide most rapidly diffused, probably due to its high water solubility, while hydrogen showed slow transport relative to the other gases. The carbon dioxide rapidly dissolved into the solution, which concurrently migrated as a clump with cations which were actively transported through the Nafion. For optimization of the reactor to maintain the purity of the

0.06

0.9

0.05

0.8 0.7

0.04

0.6 0.03 0.5 0.02

0.4

0.01

H2_Anode

0.3

0.00

CH4_Andoe CO2_Anode

0.2

CO2 (%)

3.3. Hydrogen purity and gas diffusion through Nafion membrane

H2 and CH4 (%)

whereas Rozendal et al. [3] reported the cathodic overpotential as that governing in their former study.

0.1 0

10

20

30

40

50

60

Time (hr) Fig. 6 – Gas permeability of a Nafion 117 membrane: increase of gas contents in the anode chamber of the BEC as a result of diffusion from the cathode chamber containing a standard gas mixture (4.4% CO2, 1.3% CH4 and 8.3% H2).

hydrogen produced in the cathode chamber, the exact mechanism of anodic gas diffusion should be further studied, i.e., whether the gas diffuses directly between the headspace or in the dissolved form, such as cations. Nafion was also found to be permeable to hydrogen; but interestingly, hydrogen was not detected in the anode headspace of a normally operated BEC, probably due to its instant consumption by hydrogen consuming methanogens. The high hydrogen purity is another advantage of the BEC system over other methods for producing biohydrogen from organic materials requiring additional gas purification due to diverse gaseous impurities. However, great concern is required for hydrogen contamination with the anode gases, because the Nafion membrane is severely permeable to gases produced at the anode.

3.4.

Bacterial community

Even though the analysis of the microorganism communities existing in MFCs has so far revealed a great diversity in composition [16,18–21], there has been no report on the community analysis for BEC, as a result of this research being at a premature stage. The bacterial community and the viability of anode biofilms were compared between the normal MFC and BEC, based on the concern that the artificial augmentations of the circuit by external power could generally result in changes or damage to the bacteria. The results of the 16S rDNA sequencing obtained from MFC indicted that there was great phylogenetic diversity in the communities of the anode biofilms, and was no single emergent bacterial species because electrogenesis in MFCs is due to special characteristics of bacteria but not to particular bacteria. The microbial communities in the anode biofilms of the MFC were dominated by Proteobacteria, especially the b-subclasses, which consist of several groups of aerobic or facultative bacteria (Table 1). Their predominance in MFCs was possibly due to the intrusion of oxygen from the cathode to the anode, which might support aerobic or facultative

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100

Operational mode MFC BEC

Proteobacteria (%) a

b

g

d

2.4 0.0

51.2 20.0

0.0 0.0

34.1 72.0

Firmicutes (%)

Others (%)

0.0 0.0

12.2 8.0

growth. In the acetate-enriched MFC, 51.2% of the sequences obtained in the 16S rDNA clone library obtained from an anode biofilm were b-Proteobacteria, and 66.8% of these were in the genus Thauera with a >95% similarity to Thauera aromatica LG356. The next most frequently detected bacteria was d-Proteobacteria (34.1%), with a predominance of Geobacter related species, followed by others (12.2%) and a-Proteobacteria (2.4%). For the hydrogen production, the continuous augmentations of the MFC circuit by external voltage caused remarkable changes in the bacterial community. When the MFC was switched to hydrogen producing BEC, the bacterial diversity decreased substantially with the extreme dominance of d-Proteobacteria (72% of the total bacteria) occurring (Tables 1 and 2). Most of these d-Proteobacteria were Pelobacter with a >98% similarity to Pelobacter propionicus DSM 2379, which can oxidize organic acids with Fe(III) serving as the sole electron acceptor in anaerobic sediments. The most influential electrochemically active microorganisms in the MFC systems are considered to be iron-reducing bacteria, such as Geobacter and Shewanella species, as a result of their frequent occurrence in various types of MFCs [22,23]. Geobacter-like species, which directly transfer electrons to electrodes, were determined to be integral component (16%) of the bacterial community in the BECs fed with acetate, similarly to those in the MFC. This implies an important role of Geobacter species on the exocellular electron transfer to the electrodes in the MFC-based systems. In contrast, no Shewanella-like sequences, another wellknown model strains, were retrieved from the BECs. These contrary observations of the two model species are probably attributed to the substrate they use. Geobacter species have strong affinity for acetate as an electron donor, but Shewanella species preferentially utilize lactate. Therefore, Geobacter species are more advantageous relative to Shewanella species when competing for space on the anode as acetate is used as substrate.

Live and dead cell (%)

Table 1 – Comparison of bacterial communities obtained from the MFC and BEC runs with acetate

Live cells Dead cells

80

60

40

20

0 MFC

BEC

Fig. 7 – Cell viability with respect to operational mode. Fractions of live and dead bacteria in the anode biofilms from the MFC and BEC enriched with acetate were measured using membrane integrity staining kits. Bars indicate standard deviation (n [ 10 for MFC, 26 for BEC).

3.5.

Bacterial viability

The modified MFC for the production of hydrogen appears to be twice as efficient as electricity-producing MFC, but not enough systematic studies have been conducted, especially on the anodic bacterial viability and their communities. The artificial augmentations of a circuit by external power to overcome the thermodynamic barrier resulted in a decreased cell density, but an increased cell viability (Fig. 7). The biofilm density became lower as the MFC changed to the BEC run for the production of hydrogen: the extracted amount of DNA was 78.0 mg/cm3 carbon felt for the BEC, but 535.2 mg/ cm3 for the MFC. However, the bacterial viability of the anode biofilms was enhanced from 69.0% in the MFC to 81.6% in the BEC (Fig. 7). Representative images of live and dead bacteria obtained from the MFC and the BEC are shown in Fig. 8. Such staining was mainly based on the cell membrane integrity and, therefore, does not perfectly distinguish live from dead cells, but reliably and quantitatively identify live and dead bacteria. In addition, the biofilm was observed to be more dense in the outermost few millimeters of the carbon

Table 2 – Characterization of cloned 16S rRNA gene fragments derived from an acetate-enriched BEC Isolate clone no. BEC-2, 15, 25 BEC-5, 21 BEC-1, 6–9,10–14, 20, 24 BEC-17 BEC-4 BEC-18 BEC-26 BEC-19 BEC-22

Highest homology

Occurrence (%)

Azonexus caeni Azonexus sp. RV3 Pelobacter propionicus DSM 2379 Geobacter chapelleii Geobacter sp. G02 Geobacter sp. CdA-3 Geobacter psychrophilus Uncultured bacterium RB046 Uncultured bacterium TC28

12 8 56 4 4 4 4 4 4

a Based on 727 bp of 16S rRNA sequence.

Accession no. AB166882 DQ833391 CP000482 GCU41561 EF014495 Y19191 AY653549 AB240295 EF644512

% Similarity a

Taxon

100 99 98 97 98 97 97 97 99

b-Proteobacteria b-Proteobacteria d-Proteobacteria d-Proteobacteria d-Proteobacteria d-Proteobacteria d-Proteobacteria Others Others

international journal of hydrogen energy 33 (2008) 5184–5192

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Fig. 8 – Confocal laser scanning microscopy images of live–dead bacteria from MFC (A) and BEC (B) fed with acetate. Live cells are shown in green and dead cells in red, but some areas appear yellow due to overlapping. Bars [ 1 mm. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

felt and gradually decreased with respect to depth towards the inside (data not shown). This stratification was due to the limited opportunities for contact between the substrate and the bacteria deep inside the felt as a result of the quite densely entwined structure of carbon felt, with a corresponding small void space. Therefore, sophisticated engineering of the anode will be necessary for the uniform occupation of the electricigens, both in quantity and viability, throughout the entire anode space by controlling the entwined structure of carbon felt.

the transport of protons. Hence, a number of problems associated with CEM still exist in the BEC, as with the MFC, such as diffusional hydrogen loss, cation transport rather than protons and biofouling. Furthermore, the inherent weakness of the additional voltage supply must be resolved to enhance the economic feasibility of this technology. This biocatalyzed electrolysis still at an early research stage, but with better understanding, could be a revolutionary breakthrough for capturing renewable hydrogen energy sources.

Acknowledgements 4.

Conclusion

Sufficient hydrogen gas was generated using the BEC system at applied voltages above 0.30 V and indeed originated from the degradation of acetate rather than water splitting. The hydrogen yields gradually increased with increasing applied voltage from 0.1 to 0.8 V, but decreased above 0.8 V. The anodic loss of electrons was more severe than the cathodic loss, especially for applied voltages above 0.5 V. The high purity of hydrogen produced appears an attractive advantage of the BEC system over conventional biohydrogen production methods containing diverse gaseous impurities. The bacterial communities responsible for electrogenesis differ in different operational modes (e.g., MFC and BEC) and characterization of these populations is crucial to understanding how the BEC can be controlled and optimized. There were substantially fewer bacterial species in the communities of the anode biofilms from the BEC than observed in the MFC. P. propionicus-like species were remarkably dominant in the BEC. Geobacter-like species were important species in the MFC and were still integral members of the bacterial community for the BEC. The artificial augmentations of the circuit by an external voltage did not damage the bacterial viability in the BEC. In the BEC systems, the installation of a membrane is essential to separate the anode from the cathode, but to allow

This research was supported by a grant (code# C106A152000106A085200000) from Plant Technology Advancement Program funded by Ministry of Construction & Transportation of Korean government and in part by the Center for Distributed Sensor Network at Gwangju Institute of Science and Technology (GIST).

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