Effect of static magnetic field on the performances of and anode biofilms in microbial fuel cells

Abstract

The electricity production and substrate removal rate of microbial fuel cells (MFCs) could be improved by the application of a static magnetic field (SMF). The effects of SMF intensities on cell performances and the activity of anode biofilms were investigated in this manuscript. A MFC with an intensity of 220 mT SMF obtained the best cell performances. The maximum output voltage increased from 360.1 mV to 756.1 mV and the coulombic efficiencies increased from 34.57% to 60.46% compared with the control. And the start-up time decreased from 200 h (the control) to 140 h in MFC-220. Ammonia in MFCs was removed by volatilization of ammonia, nitrification and denitrification in MFC. And nitrogen compounds in MFCs were nitrified thoroughly with SMF. With a low SMF (≤220 mT), a MFC could enhance the Coulomb efficiency by inhibiting methane production. SMF promotes power outputs by affecting the enzyme activity and bioelectrochemical activity of biofilms as well as biomass. With increased low SMF intensity, the microorganism was stimulated to produce more extracellular polymeric substance. Scanning electron microscope and confocal laser scanning microscopy images showed that bacteria and biopolymers coexisted or overlapped on the anode. However, a high intensity (≥360 mT) of the applied SMF was harmful to microbial growth and had negative effects on cell performance.

---

1. Introduction

Microbial fuel cells (MFCs) have become a promising technology in the recovery of electricity from waste by transferring electrons released from the oxidation of organics across external loading. However, the achievable power output of MFCs is still not commercially feasible. A number of attempts have been made to improve electricity generation through enhancing the oxidation of substrates, electrons flowing toward the anode, and their consumption at the cathode; such attempts include the optimization of operational parameters,¹ various modifications of electrodes,^2–4 and the design of configurations and assemblies.⁵

Previous research has indicated that the application of a static magnetic field (SMF) is a promising approach for the enhancement of biological activity.⁶ The applied SMF has an effect on the organizational structure and metabolism of living microorganisms; such an effect is called the magnetic biological effect,⁷ which involves metabolism velocity and enzyme activity.⁸ High SMF intensity inhibited the physiological processes of microbes,⁹ while lower SMF intensity motivated microbial activity. Formaldehyde biodegradation during the activated sludge process increased by 30% under 7 mT SMF and had a positive effect on microbe growth and dehydrogenase activity (DHA).¹⁰ Notably, 100 mT SMF applied on Shewanella-inoculated MFC improved the peak voltage by 20–27% in both single- and two-chamber MFCs.¹¹ The improvement in electricity production was mainly attributed to the enhanced bioelectrochemical activity, possibly through the oxidative stress mechanism.¹² SMF also affected the bioelectrocatalytic transformations of several enzyme assemblies linked to electrodes through accelerating electron transfer at the electrode–liquid interface.^13–15

Nitrogen removal is of high interest in wastewater treatment because of the tight regulations on nitrogen discharge.¹⁶ Ammonia cannot be effectively oxidized under anaerobic conditions.¹⁷ However, in a single-chamber air-cathode MFC, oxygen was allowed in the atmosphere to passively diffuse into the solution, which was possible for nitration occurred. Kim et al.¹⁸ demonstrated that the main mechanism for ammonia removal was probably not due to biological processes but rather the volatilization of ammonia at the cathode. Nitration was observed in a two-chamber MFC and was found to be more thorough with SMF.¹⁹ However, whether the application of SMF on single-chamber MFC had an effect on nitrogen removal and the mechanism for ammonia removal remains unclear.

These previous SMF studies are efficient attempts to evaluate SMF in MFCs. However, none of them analyzed effect of SMF on cell performance and biofilm change with SMF intensities of a single-chamber MFC in mixed-culture medium. In this manuscript, different intensities of SMF were applied on mixed-culture single-chamber MFC reactors. Through testing the carbon balance, nitrogen balance, nitrogen transformation, and biofilm change on/in anode, we aim to (i) determine the effect of SMF on mixed-culture MFC performance in electricity generation and substrate removal; (ii) clarify the carbon balance change in MFC under applied SMF; (iii) identify the nitrogen balance and ammonia removal mechanism of MFC under applied SMF; (iv) distinguish the characteristic change of the anode biofilm under applied SMF; and (v) observe the morphology change of anode biofilm under applied SMF.

2. Materials and methods

2.1 MFC configuration and operation

A single-chamber MFC reactor was constructed by using Lexan cylindrical chambers, each of which had a length of 4.0 cm and a diameter of 3.2 cm with a total volume of 28 mL. The anode was composed of graphite felt with a projected area of 7.1 cm² (Sanye Carbon Ltd, China). The air cathode had a projected surface area of 8.0 cm² and was made of carbon cloth (Hesen electric Ltd, China) with four PTFE diffusion layers (facing the oxygen) and a 0.5 mg_{Pt C} cm⁻² catalyst layer (facing the solution). The anode and cathode were connected by a titanium wire across an external resistor of 1000 Ω. The gap between the anode and the cathode was 3.0 cm. All reactors were operated in a sequencing batch mode at room temperature (25 °C).

The reactors were exposed to SMF intensities of 0 mT (control), 20 mT (MFC-20), 120 mT (MFC-120), 220 mT (MFC-220), and 360 mT (MFC-360), respectively. The SMF intensity was applied by binding a square magnet onto the external chamber wall near the anode side to constitute an SMF-coupled MFC. The length of the square magnets was 4.0 cm while heights differed. The tops and bottoms of the magnet were the N and S, respectively, and the top surface was near the anode surface. The magnetic induction lines across the electrode were approximately parallel to each other and perpendicular to the surface of the electrode from the anode to the cathode because the magnet was close to the electrode while the electrode was thin. SMF intensity was tested by using a teslameter (WT-10A, TES Electrical Electronic Corp, China). When the cell voltage across the external loading decreased below 50 mV for a period of approximately 2 h, the medium was refreshed to begin the next fed-batch cycle (Fig. 1).

Fig. 1 The diagrammatic drawing of MFC with SMF.

The inoculums of the MFC reactors were 12 mL effluent collected from the existing sediment MFC for sludge treating in our laboratory. The growth medium contained (per L): 1.00 g sodium acetate, 11.53 g Na₂HPO₄·12H₂O, 2.77 g NaH₂PO₄·2H₂O, 0.51 g NH₄Cl, 0.13 g KCl, 12.5 mL minerals, 5 mL vitamins,²⁰ and 0.1 g Fe(III) (provided by Fe₂(SO₄)₃·xH₂O).²¹

In order to calculate the nitrogen and carbon balance, a closed system was developed (Fig. S1†) to test the gases volumes and composes. An empty air bag was connected to cell to collect outgas, and an empty chamber was installed next to the cathode side with an oxygen bag connected to it to provide the cathode with needed oxygen.

2.2 Electrochemistry analysis

The cell voltage (V) across the external loading (R) was automatically recorded by a computer-based data acquisition system (DAQ-2204, Taiwan ADLINK Ltd, China) at a pre-determined sampling frequency (0.5 h). Coulombic efficiency (CE) was obtained by using the method introduced by Logan et al.²²

The polarization and power density curves of the SMFC reactors were obtained through a discharge test.²² Cyclic voltammetry (CV) was conducted on the anode of MFCs with the cathode as the counter electrode. A saturated calomel electrode (+0.242 V versus the standard hydrogen electrode) (Gaoshirilian Ltd, China) was used as the reference electrode by using an electrochemical workstation (CHI600D, CH Instruments Inc., China) in a fresh growth medium. Scans ranged from −0.8 V to +0.8 V at a rate of 1 mV s⁻¹ with only the fourth stable cycle shown.

2.3 Chemical analysis

The chemical oxygen demand (COD), nitrite nitrogen (NO₂⁻–N), nitrate nitrogen (NO₃⁻–N) and ammonium nitrogen (NH₄⁺–N) were analyzed according to the Chinese national environmental protection agency standard method.²³ The concentration of sulfate (SO₄²⁻) was tested by using the barium chromate colorimetric method.²⁴ Gas volumes were tested by using the drainage method. The gas composition was determined by using a gas chromatograph (GC-14C, Shimadzu, Japan) equipped with a thermal conductivity detector. The gas in both sides was absorptive into dilute sulfuric acid to test the content of NH₃ in gas.

The biomass and DHA of the anode biofilm were determined by phospholipid analysis²⁵ and the TTC-reduction method.²⁶ The soluble extracellular polymeric substances (EPS) produced by microorganisms were obtained by adding an anode electrode to 5 mL phosphate buffer saline solution (PBS, 50 mM, pH 7.0), then the mixed liquids were centrifuged at 12 000 × g (CF15RXII, Hitachi, Japan) for 10 min. The supernatant was collected and filtrated through a 0.45 μm filter membrane. The EPS was assumed the sum of extracellular proteins (PN) and polysaccharides (PS). PN were determined by using a modified Lowry method²⁷ using bovine serum albumin (Sinopharm Chemical Reagent Co., Ltd, China) as the standard. PS were measured by using the phenol–sulfuric acid method²⁸ with glucose (Sinopharm Chemical Reagent Co., Ltd, China) as the standard.

2.4 Scanning electron microscope and confocal laser scanning microscopy imaging analysis

For scanning electron microscope (SEM) analysis, part of the anode was cut into pieces and immersed in 2.5% glutaraldehyde for 30 min. The samples were carefully rinsed three times with PBS (50 mM, pH 7.0) and once with deionized water. The samples were then subjected to dehydration using an ethanol series (30%, 50%, 70%, 80%, and 95%, 30 min for each concentration) and then dried completely at ambient temperature. The microscopic structure of the anode was analyzed using SEM (SU1510, Hitachi, Japan). The composition and architecture of the biofilms on/in anodes were analyzed by confocal laser scanning microscopy (CLSM) (ZEISS LSM 710, Zeiss, Germany). Fluorescent staining was conducted as described by Chen et al.²⁹ Major fluorescence probes including Syto 63, fluorescein isothiocyanate, and calcofluor white (all purchased from Sigma Inc.), were used to simultaneously probe the cells, PN and PS, respectively.

3. Results and discussion

3.1 Electricity generation

Fig. 2 showed the change in the voltage outputs from the start to the stable operation of the MFCs exposed to different SMF intensities. From Fig. 2, we could find that the time required to reach the first maximum power cycle was significantly shorter when the MFCs were exposed to SMFs. MFC-360 with SMF of 360 mT started the fastest and required approximately 100 h before reaching maximum power production with a maximum voltage of 171.8 mV. MFC-220 followed and required approximately 140 h to reach the maximum voltage, with a highest voltage of 756.1 mV. For the control, MFC-20 and MFC-120, the times required to reach the first maximum power cycle all increased to 3 cycles approximately 200 h, and the maximum voltages were 360.1, 457.5, and 579.9 mV, respectively. These results suggested that SMF could accelerate the MFCs to stabilize. In addition, unlike the control, an obvious increase in the maximum output voltage was observed for MFCs with low SMFs except MFC-360. The maximum output voltage for MFC-360 did not increase but decreased significantly. This indicated that too high intensity would have negative effects on electricity production; otherwise SMF application was benefits to it.

Fig. 2 MFC voltage outputs change during operation period exposed to different SMF intensities.

The direction of SMF exposed to MFC was also invested in this work. During the fifth stable cycle of MFC-220, we changed the direction of SMF with the bottom surface (S) near the anode surface. And the voltage exhibited negligible difference (Fig. S2†). This was also confirmed by other researches.¹¹

3.2 Polarization and power density curves

Based on the polarization and power density curves (Fig. 3), the maximum power densities and internal resistances change with different SMF intensities were obtained. The maximum power density increased to 0.63 W m⁻² in MFC-20, to 0.88 W m⁻² in MFC-120, to 1.50 W m⁻² in MFC-220 from 0.30 W m⁻² in the control, while the maximum power density of MFC-360 only obtained 0.09 W m⁻². This again confirmed that low intensity (≤220 mT) was favorable for power generation, while high intensity (≥360 mT) was harmful to it. The activation resistance was visible in the polarization curve at low currents initially decreased with low intensities and increased with high intensity. Electron transfer from the cell terminal protein or enzyme to the electrode might cause the activation polarized internal resistance, indicating that the MFC exposed to SMF had an effect on anode biofilms. These findings indicated that the performance improvement was mainly due to the effect of SMF on the anode.

Fig. 3 Polarization and power density curves exposed to different SMF intensities.

3.3 Nitrogen balance

Nitrogen removal is of great interest in wastewater treatment.³⁰ But nitrogen balance was barely investigated. In this study, we investigated the nitrogen balance and the ammonia removal mechanism with different SMF intensities in MFCs.

The nitrogen balance was divided into four parts: the residual ammonia nitrogen (NH₄⁺–N), nitrogen oxide in liquid (NO_X–N), ammonia in gas phase (NH₃) and the others (Fig. 4). From Fig. 4, we could found that with low SMF intensities (≤220 mT) the removal of NH₄⁺–N was increased while with high intensity (≥360 mT) the removal was declined compared with the control. The NO_X–N represented the part of nitrification in MFC. With low SMF intensities, the proportion of NO_X–N in total nitrogen was around 22% showed a slight increase compared to the control (18.5%). And the proportions increased a little with low SMF intensities. With high SMF intensity, the proportion of NO_X–N in total nitrogen decreased to 7.8%. This indicated that in low SMF-coupled MFC the nitrification was slightly enhanced while with high SMF intensities the nitrification was inhibited. The proportion of NH₃ in all MFCs was around 27% indicated that the volatilization of ammonia was not influenced by SMF. In addition to the above three parts, there was some nitrogen unaccounted for by these tests and defined as the others. By tested the gas compose (shown in Table S1†), there might have a very small amount of N₂ appeared in the gas phase, the possible analytic of this part might be due to the denitrification on/in the anode. And this part showed similar rule with NO_X–N change.

Fig. 4 Nitrogen balance of MFCs exposed to different SMF intensities.

The concentrations of NH₄⁺–N, NO₃⁻–N, and NO₂⁻–N before and after operation were measured. Results are shown in Table 1. Ammonium decreased, and nitrate and nitrite were produced and accumulated after operation, thereby indicating the occurrence of nitration. Ammonia was detected in gas phase indicated that part of the NH₄⁺–N was removed by volatilization. Nitrogen compound transformations in the SMF-coupled MFCs and the control were significantly different. In the control, a large proportion of NH₄⁺–N was converted into NO₂⁻–N; however, more NH₄⁺–N was converted into NO₃⁻–N in SMF-coupled MFCs. Nitrification is a two-step process: NH₄⁺ is first oxidized into NO₂⁻, and then further oxidized into NO₃⁻. The possibility of partial nitrification occurred in the single-chamber MFC cathode but more thorough nitrification with SMF does exist; this finding is consistent with previous research.^19,31 In MFCs, the ammonium removal mechanism was divided into two parts: one was ammonia volatilization and the other was nitrification and denitrification during operation.

Table 1 NH₄⁺–N, NO₃⁻–N and NO₂⁻–N concentrations in MFCs at the end of the cycles exposed to different SMF intensities

	NH4^+–N (mg L^−1)	NO3^−–N (mg L^−1)	NO2^−–N (mg L^−1)
Initial	133.4	0	0
Control	61.8 ± 1.2	1.7 ± 0.3	23.0 ± 1.7
MFC-20	56.7 ± 1.8	16.1 ± 0.6	10.8 ± 0.3
MFC-120	54.4 ± 2.1	18.5 ± 1.7	10.2 ± 0.2
MFC-220	48.5 ± 1.1	21.8 ± 2.7	9.5 ± 0.4
MFC-360	84.6 ± 0.5	6.7 ± 0.5	3.8 ± 0.4

---

3.4 Carbon balance

Three main processes may occur in the anode: electricity production process (1), acetic fermentation to produce methane (2), and sulfate reduction (3).¹⁶

CH₃COO⁻ + 4H₂O → 2HCO₃⁻ + 9H⁺ + 8e⁻ (1)

CH₃COO⁻ + H₂O → CO₂ + CH₄ + OH⁻ (2)

CH₃COO⁻ + SO₄²⁻ + H₂O → 2CO₂ + 2H₂O + S²⁻ + OH⁻ (3)

A mass balance of carbon compounds based on COD was established with the MFCs by analyzing the contributions from different sources, including electricity, methane, sulfate and other unknown factors (Fig. 5). This balance could also represent an electron balance because carbon is an electron donor. The carbon distribution to electricity production was derived from CE with 34.57 ± 1.20%, 43.08 ± 0.75%, 52.78 ± 1.06%, 60.46 ± 1.46%, and 29.80 ± 3.00% for the control, MFC-20, MFC-120, MFC-220 and MFC-360, respectively. Eqn (2) shows that 64 g of COD is consumed to produce 1 mol of methane. Gas volume and composition of the air bag was shown in ESI (Table S1†). And in the oxygen bag, we had not detected other composition except oxygen by gas chromatography. The proportion of oxygen was all above 98% in all oxygen bags. The methane production was measured in gas phase with 2.29 ± 0.04, 2.15 ± 0.10, 1.31 ± 0.05, 0.98 ± 0.06, and 0.41 ± 0.01 mL. The number of moles of CH₄ was calculated by the ideal gas law (PV = nRT, assuming the actions occurred under a standard atmospheric pressure with 25 °C). Calculation results indicate that the methane productions consumed carbon of 34.48 ± 0.74%, 30.86 ± 1.97%, 17.01 ± 1.00%, 13.87 ± 0.61%, and 8.13 ± 0.58% COD for the control, MFC-20, MFC-120, MFC-220, and MFC-360, respectively. Eqn (3) indicates that 1 g SO₄²⁻ removal needs 0.67 g COD. The fresh growth medium contained a sulfate concentration of 257 mg_SO4²⁻ L⁻¹, and the anodes removed 81.12 ± 0.61%, 82.23 ± 0.60%, 84.01 ± 0.75%, 84.18 ± 0.80%, and 74.60 ± 0.33% sulfate, thereby resulting in 22.54 ± 0.58%, 21.73 ± 0.47%, 22.32 ± 0.84%, 21.85 ± 0.53%, and 38.47 ± 5.13% of carbon being consumed by the control, MFC-20, MFC-120, MFC-220, and MFC-360, respectively. A sizable COD was still unaccounted for by these calculations, thereby indicating that another process occurred aside from the three main processes in MFCs. And a small amount of hydrogen production process may occur in MFCs (Table S1†).³² The carbon balance indicated that SMF could effectively inhibit acetic fermentation to produce methane but barely had an effect on sulfate reduction. However, sulfate reduction consumes a considerable amount of carbon. Biological sulfate reduction should be inhibited to enhance the electron recovery efficiency.

Fig. 5 Carbon balance based on COD in MFCs exposed to different SMF intensities.

3.5 Effect of SMF intensities on microbial growth

To obtain a thorough insight into the effect of SMF intensities on electricity generation, the microbial growth changes derived under different SMF intensities were determined and showed in Table 2. The morphology change of microbial growth was tested by using SEM (Fig. 6). The SEM images showed that the anode consisted of microorganisms with EPS. And with low SMF intensities, more microorganisms appeared to attach on the carbon fiber. In addition, microorganism distribution became compact, with more EPS attached on the carbon fiber with low SMF intensities; however, almost no obvious biofilm appeared in MFC-360, thereby indicating that high intensity harmed biofilm growth. The microorganisms attached to the carbon fiber had a uniform morphology, exhibiting a rod type species primarily.

Table 2 Anode biofilm change of MFCs under different SMF intensities

	Control	MFC-20	MFC-120	MFC-220	MFC-360
Biomass (μgP cm^−3)	54.66 ± 0.52	57.65 ± 0.57	58.07 ± 0.57	60.31 ± 0.67	38.75 ± 0.79
TTC-DHA (μgTF (cm^3 h)^−1)	50.87 ± 0.31	109.83 ± 0.34	134.30 ± 0.53	147.50 ± 0.54	48.39 ± 0.68
TTC-DHA (μgTF (μgP h)^−1)	0.93 ± 0.00	1.91 ± 0.01	2.31 ± 0.01	2.45 ± 0.02	1.24± 0.01
PN (μg cm^−3)	14.05 ± 0.25	16.32 ± 0.72	16.88 ± 0.88	20.49 ± 0.57	13.43 ± 0.59
PS (μg cm^−3)	15.41 ± 0.53	19.58 ± 0.55	24.79 ± 0.62	28.36 ± 0.61	13.14 ± 0.76
EPS (μg cm^−3)	29.46 ± 0.77	35.90 ± 1.28	41.67 ± 1.49	48.86 ± 1.17	26.56 ± 0.61
EPS/biomass (μg/μgP)	0.54 ± 0.01	0.63 ± 0.02	0.72 ± 0.02	0.81 ± 0.01	0.69 ± 0.02

---

Fig. 6 SEM images of carbon felt (A) and anodes of the control (B), MFC-20 (C), MFC-120 (D), MFC-220 (E) and MFC-360 (F) with different SMF intensities.

We monitored the phospholipid on the anodes to determine the biomass. We found that the biomass on/in the anode biofilms changed significantly with low SMF intensities; the obtained biomass that exposed to SMF was raised with SMF intensities, while MFC-360 obtained the least amount of biomass with 38.75 μg_P cm⁻³. With low SMF intensities the biomass of MFCs were increased. While with high SMF intensity the biomass showed a significant decline. This indicated that high SMF intensity was harmful to microorganisms. This was also verified by SEM images.

The promotion of power output with low SMF intensities may also due to the enhancement of enzyme activity and bioelectrochemical activity of microorganisms.³³ As expected, the TTC-DHA activity first increased with low SMF intensities and then decreased with high SMF intensity (Table 2); this finding was similar to reported findings.^14,15 The highest TTC-DHA activity of 147.50 ± 0.54 μg_TF (m³ h)⁻¹ was achieved in MFC-220 and decreased significantly to 48.39 ± 0.68 μg_TF (m³ h)⁻¹ in MFC-360, which was even lower than the control at 50.87 ± 0.31 μg_TF (m³ h)⁻¹. Considering the biomass change in anodes, we calculated the TTC-DHA change per mg_P biomass (Table 2). The results showed that low SMF intensities could great boost the microbial activity. However, unlike the above results, the TTC-DHA activity per mg_P biomass remained higher than the control in MFC-360. This indicated that the restraining electricity generation in MFC-360 was mainly due to the highly inhibited of bacterial growth but not to the effect of microbial activity compared with the control.

CVs were performed to examine the bioelectrochemical activity of biofilms with different SMF intensities (Fig. 7). All biofilms showed bioelectrocatalytic activity, and the maximum and minimum currents changed with SMF intensities. The maximum and minimum peak currents varied for biofilms and acclimated with different SMF intensities with values of 0.038 and −0.12 mA (the control), 0.073 and −0.14 mA (MFC-20), 0.079 and −0.25 mA (MFC-120), 0.11 and −0.34 mA (MFC-220), 0.037 and −0.041 mA (MFC-360), respectively. The biofilm with 220 mT showed the highest bioelectrochemical activity, followed by biofilms with 120 mT, 20 mT, the control, and 360 mT; this finding was consistent with the maximum currents produced in MFCs. No diffusional current decay should occur with fresh growth medium. Therefore, the different steady-state currents could likely reflect the transport of redox species through these biofilms.

Fig. 7 Cyclic voltammograms of the biofilms of MFCs exposed to different SMF intensities in fresh growth medium with 1 g L⁻¹ sodium acetate.

3.6 EPS change with SMF

EPS was beneficial to microbial growth mainly because it protected microbes from negative environmental variations.³⁴ EPS existed in the complex and heterogeneous polymeric matrix, and played a vital role in maintaining the stable structure of biofilms. PN and PS were the two main constituents of EPS. The changes in PN, PS and EPS were shown in Table 2. The MFC-220 reactor achieved the highest levels of PN, PS, and EPS with 20.49, 28.36, and 48.86 μg cm⁻³, respectively, which was consistent with the trend of anode biomass. The PS and PN contents in anode biofilm with low SMF intensities increased. SMF could stimulate microbial growth to produce EPS, and the content was enhanced with low SMF intensities. MFC-360 obtained the lowest levels of PN (13.43 μg cm⁻³), PS (13.14 μg cm⁻³) and EPS (26.56 μg cm⁻³), which were even lower than the control. We calculated the production of EPS per mg_P biomass. Similar to the process of calculating TTC-DHA, the content of EPS per mg_P biomass was higher compared with the control. The slight decrease of EPS content in MFC-360 was due to the damage caused by high SMF on microbial growth.

Aside from microbial EPS, we used CLSM images for fixed biomass, PS, and PN (Fig. 8). Bacteria (red), β-polysaccharides (green), and proteins (blue) were present on the anodes. High contents in the anode corresponded to brighter images. The images indicated that both bacteria and biopolymers coexisted or overlapped on/in the anode. Biopolymers can be adsorbed easily onto the anode surface and adhere on bacterial cells, which might facilitate initial microorganism deposition. The images also showed that PS, PN, and microbial cells were aggregated into clusters. The experimental groups with SMFs have the same morphology; this finding is similar to findings in other research.^35,36 Similar to the content test, the different fluorescence intensities of CLSM images showed that the PN and PS in anodes changed with SMF intensities; the contents of bacteria, PS and PN first increased then decreased with SMF intensities; this trend is the same as that of power generation.

Fig. 8 CLSM images of biofilm on/in the anodes of the control (A), MFC-20 (B), MFC-120 (C), MFC-220 (D), MFC-360 (E) of MFCs exposed to different SMF intensities (control, 20, 120, 220, and 360 mT), respectively (left to right in all lines-CLSM image of microbial cells (Syto 63) denoted in red; CLSM image of proteins (stained with FITC) denoted in green; CLSM image of β-polysaccharides (calcofluor white) denoted in blue and CLSM image of combined image of individual images).

4. Conclusion

Low SMF intensity was highly efficient in enhancing the electricity generation of mixed-culture MFC because of its unique feature, that is, the ability to derive high-activity biofilm in the anode.

The cell performance of MFC improved with the increase of low SMF intensities (≤220 mT). The MFC obtained the best output voltage and pollutant removal with the application of 220 mT SMF in this study. However, high intensities (≥360 mT) of the applied SMF had negative effects on cell performance.

Nitrogen balance indicated that in MFCs 27% ammonia removed by volatilization of ammonia which was not influenced by SMF, and the residual ammonia was removed by nitrification and denitrification which was influenced by SMF intensities. The possibility of partial nitrification and denitrification occurred in MFCs. More thorough nitrification existed with the application of SMF than in the control.

The carbon balance indicated that the undesired reaction of methane fermentation was restrained, and more energy was converted into electric energy with SMF intensity. Biological sulfate removal rate was above 74% in all MFCs which was not affect by SMF. Biological sulfate reduction taken a considerable proportion of total carbon and should be further inhibited to enhance the electron recovery efficiency.

SMF intensity could efficiently improve the bioelectrochemical activity and TTC-DHA activity of microbes as while as enhance the biomass. Microbial growth was stimulated to produce more EPS with different SMF intensities. Moreover, rod-type bacteria and biopolymers simultaneously coexisted or overlapped on the anode surface.

Acknowledgements

This work is supported by a grant from the Major Science and Technology Program for Water Pollution Control and Treatment of China (No. 2012ZX07101-013-04) and the Fundamental Research Funds for the Central Universities (JUSRP51512).

References

1. T. S. Song, Z. S. Yan, Z. W. Zhao and H. L. Jiang, J. Chem. Technol. Biotechnol., 2010, 85, 1489–1493.
2. D. Hidalgo, T. Tommasi, V. Cauda, S. Porro, A. Chiodoni, K. Bejtka and B. Ruggeri, Energy, 2014, 71, 615–623.
3. Y. P. Ren, D. Y. Pan, X. F. Li, F. Fu, Y. N. Zhao and X. H. Wang, J. Chem. Technol. Biotechnol., 2013, 88, 1946–1950.
4. L. J. Ren, J. C. Tokash, J. M. Regan and B. E. Logan, Int. J. Hydrogen Energy, 2012, 37, 16943–16950.
5. G. Papaharalabos, J. Greenman, C. Melhuish and I. Ieropoulos, Int. J. Hydrogen Energy, 2015, 40, 4263–4268.
6. N. S. Zaidi, J. Sohaili, K. Muda and M. Sillanpää, Sep. Purif. Rev., 2014, 43, 206–240.
7. R. L. Moore, Can. J. Microbiol., 1979, 25, 1145–1151.
8. H. Sahebjamei, P. Abdolmaleki and F. Ghanati, Bioelectromagnetics, 2007, 28, 42–47.
9. J. Miyakoshi, Prog. Biophys. Mol. Biol., 2005, 87, 213–223.
10. M. Łebkowska, A. Rutkowska-Narożniak, E. Pajor and Z. Pochanke, Bioresour. Technol., 2011, 102, 8777–8782.
11. W. W. Li, G. P. Sheng, X. W. Liu, P. J. Cai, M. Sun, X. Xiao, Y. K. Wang, Z. H. Tong, F. Dong and H. Q. Yu, Biosens. Bioelectron., 2011, 26, 3987–3992.
12. Y. Yin, G. T. Huang, Y. R. Tong, Y. D. Liu and L. H. Zhang, J. Power Sources, 2013, 237, 58–63.
13. E. Katz, O. Lioubashevski and I. Willner, J. Am. Chem. Soc., 2004, 126, 11088–11092.
14. C. Niu, J. J. Geng, H. Q. Ren, L. L. Ding, K. Xu and W. H. Liang, Bioresour. Technol., 2013, 150, 156–162.
15. L. O. Santos, R. M. Alegre, C. Garcia-Diego and J. Cuellar, Process Biochem., 2010, 45, 1362–1367.
16. F. Zhang, Z. Ge, J. Grimaud, J. Hurst and Z. He, Environ. Sci. Technol., 2013, 47, 4941–4948.
17. Z. He, J. J. Kan, Y. B. Wang, Y. L. Huang, F. Mansfeld and K. H. Nealson, Environ. Sci. Technol., 2009, 43, 3391–3397.
18. J. R. Kim, Y. Zuo, J. M. Regan and B. E. Logan, Biotechnol. Bioeng., 2008, 99, 1120–1127.
19. Q. Q. Tao and S. Q. Zhou, Appl. Microbiol. Biotechnol., 2014, 98, 9879–9887.
20. S. A. Cheng, D. F. Xing, D. F. Call and B. E. Logan, Environ. Sci. Technol., 2009, 43, 3953–3958.
21. Y. N. Zhao, X. F. Li, Y. P. Ren and X. H. Wang, RSC Adv., 2016, 6, 47974–47980.
22. B. E. Logan, B. Hamelers, R. A. Rozendal, U. Schröder, J. Keller, S. Freguia, P. Aelterman, W. Verstraete and K. Rabaey, Environ. Sci. Technol., 2006, 40, 5181–5192.
23. Chinese N. Water and Wastewater Monitoring Methods, Chinese Environmental Science Publishing House, Beijing, 4th edn, 2002.
24. H. L. Golterman and I. M. D. Bierbrauwerwurtz, Hydrobiologia, 1992, 228, 111–115.
25. R. H. Findlay, G. M. King and L. Watling, Appl. Environ. Microbiol., 1989, 55, 2888–2893.
26. H. V. Bergmeyer, W. R. Heney and J. Clin, Pathology, 1987, 2, 934.
27. B. Frølund, R. Palmgren, K. Keiding and P. H. Nielsen, Water Res., 1996, 30, 1749–1758.
28. M. Dubois, K. A. Gilles, J. K. Hamilton, P. T. Rebers and F. Smith, Anal. Chem., 1956, 28, 350–356.
29. M. Y. Chen, D. J. Lee, J. H. Tay and K. Y. Show, Appl. Microbiol. Biotechnol., 2007, 75, 467–474.
30. N. Lu, S. G. Zhou, L. Zhuang, J. T. Zhang and J. R. Ni, Biochem. Eng. J., 2009, 43, 246–251.
31. A. Tomska and L. Wolny, Desalination, 2008, 222, 368–373.
32. J. C. Tokash and B. E. Logan, Int. J. Hydrogen Energy, 2011, 36, 9439–9445.
33. N. S. Zaidi, J. Sohaili, K. Muda and M. Sillanpää, Sep. Purif. Rev., 2014, 43, 79–92.
34. G. P. Sheng, H. Q. Yu and X. Y. Li, Extracellular polymeric substances (EPS) of microbial aggregates in biological wastewater treatment systems: a review, Biotechnol. Adv., 2010, 28, 882–894.
35. Y. K. Wang, G. P. Sheng, W. W. Li, Y. X. Huang, Y. Y. Yu, R. J. Zeng and H. Q. Yu, Environ. Sci. Technol., 2011, 45, 9256–9261.
36. J. Xu, G. P. Sheng, H. W. Luo, W. W. Li, L. F. Wang and H. Q. Yu, Water Res., 2012, 46, 1817–1824.

---

Footnote

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c6ra15844g

---