Electricity and biomass production in a bacteria-Chlorella based microbial fuel cell treating wastewater

Journal of Power Sources xxx (2017) 1e11

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Electricity and biomass production in a bacteria-Chlorella based microbial fuel cell treating wastewater Audrey S. Commault*, Olivier Laczka, Nachshon Siboni, Bojan Tamburic, Joseph R. Crosswell, Justin R. Seymour, Peter J. Ralph Climate Change Cluster (C3), University of Technology Sydney, Ultimo, NSW, 2007, Australia

h i g h l i g h t s
Diurnal cycle of the power output with Chlorella vulgaris at the cathode.
Maximum power doubled to 34.2 mW m2 in the presence of C. vulgaris.
The majority (90%) of C. vulgaris cells were attached to the cathode surface.
0.19 gL1d1 of chemical oxygen demand and 5 mgL1d1 of ammonium was removed.
Cathodic algal-bacterial community dominated by nitrogen-cycling bacteria.

a r t i c l e i n f o

a b s t r a c t

Article history: Received 11 January 2017 Received in revised form 20 March 2017 Accepted 21 March 2017 Available online xxx

The chlorophyte microalga Chlorella vulgaris has been exploited within bioindustrial settings to treat wastewater and produce oxygen at the cathode of microbial fuel cells (MFCs), thereby accumulating algal biomass and producing electricity. We aimed to couple these capacities by growing C. vulgaris at the cathode of MFCs in wastewater previously treated by anodic bacteria. The bioelectrochemical performance of the MFCs was investigated with different catholytes including phosphate buffer and anode effluent, either in the presence or absence of C. vulgaris. The power output fluctuated diurnally in the presence of the alga. The maximum power when C. vulgaris was present reached 34.2 ± 10.0 mW m2, double that observed without the alga (15.6 ± 9.7 mW m2), with a relaxation of 0.19 gL1 d1 chemical oxygen demand and 5 mg L1 d1 ammonium also removed. The microbial community associated with the algal biofilm included nitrogen-fixing (Rhizobiaceae), denitrifying (Pseudomonas stutzeri and Thauera sp., from Pseudomonadales and Rhodocyclales orders, respectively), and nitrate-reducing bacteria (Rheinheimera sp. from the Alteromonadales), all of which likely contributed to nitrogen cycling processes at the cathode. This paper highlights the importance of coupling microbial community screening to electrochemical and chemical analyses to better understand the processes involved in photo-cathode MFCs. © 2017 Elsevier B.V. All rights reserved.

Keywords: Microbial fuel cells Chlorella vulgaris Wastewater treatment Chlorella-associated bacterial community Nitrogen cycle Algae-bacteria interaction

1. Introduction In socio-economic situations that advocate sustainability and resource management, waste repurposing is a fundamental component. It is in this context that research centred on resource recovery from wastewater has expanded, with a focus on microalgae biomass production and electricity [1,2]. Recycling wastewater reduces treatment plant operational costs and pressure on

* Corresponding author.. E-mail address: [email protected] (A.S. Commault).

valuable resources such as clean water. Microbial fuel cell (MFC) is one of the technologies used to generate electricity from wastewater. MFCs produce electricity from the anaerobic oxidation of organic matter present in wastewater by electroactive bacteria. These systems comprise of an anode and a cathode chamber connected together via a cation exchange membrane (CEM). The electrodes are linked by an external resistor allowing for electrons to flow from the anode to the cathode, creating a current. The protons generated at the anode by the anaerobic degradation of organic matter diffuse through the CEM to the cathode where they

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react with oxygen and electrons to form water. Oxygen is the most commonly used electron acceptor in MFCs due to its abundant availability and high redox potential [3]. The generation of electricity by MFCs using oxygen as an electron acceptor is hampered by two major bottlenecks: (i) the slow rate of oxygen reduction on the surface of graphite/carbon electrodes that leads to a high cathodic over-potential [4,5] and (ii) the formation of a pH gradient, from acidic conditions at the anode to alkaline conditions at the cathode, caused by the slow diffusion of protons through the CEM in competition with other cations [6], consequently reducing the availability of protons at the cathode and can damage the anodic biofilm [7,8]. The cathodic over-potential for oxygen can be lowered by the use of mediators [5], and/or expensive metal-based catalysts such as platinum [9], or biocatalysts [10,11]. The latter rely on microorganisms that have the capacity to catalyse the reduction of oxygen using electrons delivered by the cathode, as described by Bergel et al. [11], with a seawater biofilm and a stainless-steel cathode. To limit the formation of a pH gradient, some researchers have created “membrane-less” MFCs where the CEM has been completely removed [12]. However, the absence of the CEM allows for oxygen to diffuse from the cathode to the anode, where it competes with the electrode as an electron acceptor, leading to a low coulombic efficiency. To overcome the limitation caused by the creation of a pH gradient, Freguia et al. [13] introduced a new configuration in which the effluent of an acetate-fed anode was used as a feed for an aerated, biocatalysed cathode. This strategy provides an additional path for the transport of protons from the anode to the cathode. It also transfers the remaining organics from the anode compartment to the aerated cathode compartment for further processing [13]. The main disadvantage of this configuration is that it requires daily monitoring to avoid overloading the cathode with organic matter, which can lead to a considerable increase in the growth of aerobic heterotrophs. This is problematic, because aerobic heterotrophs reduce oxygen, potentially restricting the oxygen supply at the cathode and thereby, impairing electricity generation [13]. This MFC configuration also requires the aeration of the catholyte in order to provide oxygen for the cathodic reaction [13]. To counter these obstacles, this study introduced the photosynthetic microalga Chlorella vulgaris at the cathode, so that it would supply oxygen to the cathodic reaction [14]. C. vulgaris is known to be naturally present in wastewater [15,16] and tolerant of high loads of organic pollutants [17]. C. vulgaris is able to readily assimilate phosphorus and nitrogen from wastewater [18], making it a suitable organism for both the removal of nutrients and for biomass production. Numerous studies have demonstrated the efficiency of C. vulgaris in treating a variety of wastewaters from the dairy [19], textile [20] and swine industries [21], as well as tertiary municipal wastewater [22]. Notably, other studies have shown that growing C. vulgaris in optimized culture medium (e.g. Bold's basal medium) at the cathode of MFCs can highly enhance the power output [23e26]. The current study is based on the novel idea that anodic effluent is likely to still contain large amounts of ammonium, nitrate, phosphate, and dissolved inorganic carbon [27,28]. These nutrients are beneficial for algal growth and can enhance algal biomass production. The assimilation of these nutrients into algal biomass contributes to the cleaning of wastewater. Therefore, instead of using an optimized culture medium, as commonly employed in MFCs growing C. vulgaris, we investigated the potential of growing the alga in the cathode chamber of an MFC, using wastewater that was previously treated in the anode chamber. Specifically, the aim of this work was to understand the effect of C. vulgaris on the power output, nutrient removal and composition of the cathodic bacterial community. To address these objectives, the bioelectrochemical performance of the MFC and the removal of chemical oxygen

demand (COD), nitrate, and ammonium were measured in the presence and in the absence of C. vulgaris at the cathode. This novel design prevents the formation of a pH gradient and the use of expensive cathode catalysts and catholytes while combining biomass and electricity production with wastewater treatment. The study goes beyond what have previously been done, as treating anodic effluent at the cathode of MFCs using C. vulgaris has not being tested before. 2. Methods 2.1. Photo-cathode MFC set-up and operation The H-shaped photo-cathode MFCs (two replicates ran in parallel: MFC1, MFC2) were composed of 100 mL anodic and cathodic chambers connected together via 4.9 cm2 of cation-exchange membrane (Ultrex CMI-7000, Membranes International Inc., USA) (Fig. 1a). The cation exchange membrane was pre-conditioned before use by immersion in 5% NaCl solution at 40  C for 24 h to allow for membrane hydration and expansion. The anode and cathode were made of graphite plates (Graphtek LLC, USA) with dimensions 4  1  0.32 cm (total surface area of 0.00112 m2) and 4  2  0.32 cm (total surface area of 0.00198 m2), respectively. The graphite plates were chosen for their flat surface to facilitate the recovery of the biofilms for microbial community analysis. The graphite plate electrodes were polished with fine sandpaper and rinsed with distilled water before use. For the enrichment of the anodic biofilm, the anode and the cathode were first connected through a potentiostat (Electrochemical analyzer, CH Instruments; or mAutolab type 3, Metrohm) along with a reference electrode (Ag/ AgCl, 1 M KCl) for two months. The anode chamber was inoculated with a 50: 50 mixture of ethanol medium and freshwater sediment collected in Busbys Pond (33.897871E, 151.228508 N, Sydney, Australia). The ethanol medium was prepared with 0.1 M phosphate buffer as Na2HPO4/ KH2PO4 (pH ¼ 7.5), 7.1 mM NH4Cl, 15 mM absolute ethanol (molecular grade), and 3 mM Sodium 2-bromoethanesulfonate (BES) 98% (Sigma) to inhibit methanogenesis, autoclaved and then supplemented with a filter-sterilized mineral solution as previously described [29]. The anodic biofilm was enriched in electroactive bacteria by maintaining the anode at a potential of 0.36 V vs Ag/ AgCl (1 M KCl) while feeding it with 50 mL of fresh ethanol medium every 3e5 days to encourage growth. The medium was sparged for 10 min with nitrogen gas before feeding to ensure anaerobic conditions inside the chamber. After 41 days, the ethanol medium was replaced with synthetic wastewater, composed of (per litre of tap water): 16 g of peptone, 11 g of meat extract, 3 g of urea, 2.8 g K2HPO4, 0.2 g of MgSO4$7H2O, 0.4 g CaCl2$2H2O, and 0.7 g of NaCl. The synthetic wastewater was diluted 25-fold (referred to as SWW4% in the text) in 0.1 M phosphate buffer (as Na2HPO4/ KH2PO4). The SWW4% had a conductivity of 13 mS cm1, corresponding to a salinity of 7.9 g L1 (Multi 3430, WTW, Germany), a COD of 2922 ± 66 mg L1, 135 ± 1 mg L1 of ammonium, 165 ± 24 mg L1 of nitrate, and 9.5 ± 0.4 g L1 of phosphate. After two months of enrichment, the anodic biofilm was established and the generated current was stable. The anode and cathode were then disconnected from the potentiostat and connected via a 1 kU resistor, thereby maintaining a low anode potential between 0.51 V and 0.46 V vs Ag/AgCl. In theory, a low anode potential should keep the structure of the bacterial anodic community stable. The surface of the cathode was doubled (two clean electrodes of 0.00198 m2) to favour the reduction of oxygen. The cathode was originally filled with 0.1 M phosphate buffer (as Na2HPO4/KH2PO4) for the duration of the enrichment phase and the first 15 days of the 1 kU-configuration. Then, the catholyte was

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Fig. 1. (a) Photo-cathode MFC set-up schematic, (b) and cathodic biofilms in presence of C. vulgaris (top: MFC1, bottom: MFC2). After two months of anodic biofilm enrichment, the anode was disconnected from the potentiostat and connected to the cathode through a 1 kU resistor to evaluate the performance of the system with a variety of catholytes: phosphate buffer, anodic effluent (4 days old) with and without C. vulgaris. The anode-treated wastewater was used as algal nutrient feed at the cathode.

replaced by 4 day-old anodic effluents for 5 days before addition of C. vulgaris. The anodic effluent contained phosphate buffer, to ensure valid comparison of the three different catholytes tested, the same concentration of buffer was used in all of them. To maintain a high current production, the anolyte and the catholyte were replenished with fresh SWW4% and “fresh” anodic effluents every 3e5 days, respectively. The catholyte was constantly sparged with air at a flow rate of 31 cm3 min1 to supply the cathode with oxygen in both presence and absence of the alga. The cathode chamber was thoroughly cleaned with 70% ethanol between experiments with each catholyte (e.g. phosphate buffer, anodic effluent without algae, and anodic effluent with algae) to avoid cross-contamination. C. vulgaris (strain CS-42, Australian National Algae Culture Collection) was pre-grown in 50 mL of filter-sterilized anodic effluents in a shake flask at 25  C and under a light intensity of 50 mmol photons m2 s1 for 5 days. The algal culture was then used to inoculate the catholyte at a final cell density of 1  106 cells mL1 (OD750nm of 0.1), dilution (1:10). The MFC was operated at room temperature (24 ± 0.5  C); the anode chamber was kept in the dark (to prevent the production of oxygen by photosynthetic organisms), while the cathode chamber was illuminated with cool white light at an intensity of 200 mmol m2 s1 with a 12: 12 h light cycle (Fig. 1). The voltage delivered by the system was recorded every 10 s by the potentiostat. Chlorophyll a fluorescence, pH, dissolved oxygen, nutrients, and electrochemical efficiency (power curve, cyclic voltammetry) were regularly measured. 2.2. Chlorophyll a fluorescence measurement Pulse-amplitude modulated fluorometry (PAM) was used to measure the maximum quantum yield of photosystem II (Fv/Fm calculated as Fv ¼ Fm  Fo) for rapid assessment of C. vulgaris photosynthetic activity during the experiment, and to monitor for any photosystem stress caused by MFC conditions. The catholyte was stirred to bring the algae back to suspension and measurements were made with a POCKET-PAM (Gademann Instruments GmbH, Germany) at 17:00 (end of the light period), after 10 min of

dark acclimation, against the outside of the cathode chamber at approximately 1/3 height. These measurements were performed at room temperature with the following settings: blue light, measuring light intensity <0.2 mmol photons m2 s1 PAR, saturation pulse intensity of 2700 mmol photons m2 s1 PAR, and saturation pulse width of 0.6 s. 2.3. Electrochemical analyses The bioelectrochemical performance of the MFC was assessed by power curves and cyclic voltammetry. The power curves were recorded under substrate-saturated conditions using the potentiostat and a two-electrode cell configuration by coupling the Ag/ AgCl (1 M KCl) reference electrode with the counter electrode (clean graphite plate of 0.00396 m2) and applying a voltage to the anode against the counter/reference electrode, as described by Picot et al. [30]. Ten different voltages were applied over 300 s from open circuit potential to near-short circuit potential, while monitoring the steady state current. In the presence of C. vulgaris, the power curves were performed at maximum oxygen concentration. The maximum power densities (Pmax) were calculated as: Power density (W m2) ¼ (U  I) ÷ S.A., where I is the current delivered by the system at the voltage U applied, and S.A. the surface area of the anode of 0.00112 m2. To calculate the volumetric power density (W m3), the S.A. was replaced by the anode volume (0.0001 m3). The internal resistance (Rint) of the system is the resistance calculated at maximum power density using Ohm's law. Cyclic voltammetry was performed using the potentiostats at 1 mV s1 in turnover conditions (fresh SWW4% for the anode or maximal dissolved oxygen concentration for the cathode) at potentials ranging from 0.8 V to 0.2 V and 1 V to 1 V vs Ag/AgCl for the anode and cathode, respectively. 2.4. Chemical oxygen demand and N analyses To evaluate COD and N removal, samples were taken after a 4day incubation of the anodic biofilm in SWW4%, and the cathodic

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biofilm in 4 day-old anodic effluent with and without C. vulgaris. The 10 mL samples were filter-sterilized (0.22 mm Minisart, Sartorius) and stored at 20  C prior to analysis. The COD was estimated using a close reflux colorimetric method involving the addition of 2 mL samples to High Range COD Digestion vials (20e1500 mg L1, HACH Pacific, Australia). The vials were heated at 150  C for 2 h and absorbance was read against a blank with a spectrophotometer (HACH DR2010; method 435). The amount of COD removed by the biofilms was then determined as the difference between the measured COD after 4 days of incubation and the initial COD of the corresponding medium. The Coulombic efficiency was estimated as previously explained by Commault et al. [29]. The concentration of ammonium in the samples was determined using a colorimetric method, where ammonium reacted with hypochlorite ions generated by the alkaline hydrolysis of sodium dichloroisocyanurate to form monochloramine. Monochloramine then reacted with salicylate ions in the presence of sodium nitroprusside at around pH 12.6 to form a blue compound. The absorbance of this compound was measured spectrophotometrically at a wavelength of 660 nm and was related to the ammonium concentration by means of a calibration curve. The entire analysis was performed automatically using an automated photometric analyzer (Gallery™ Plus; Thermo Fisher Scientific). The measurement of nitrate was carried out using an ion chromatograph (Metrohm; model 790 Personal IC) equipped with an auto sampler and conductivity cell detector. Na2CO3 (3.2 mmol L1) and NaHCO3 (1.0 mmol L1) were used as a mobile phase with a flow rate of 0.7 mL min1. 2.5. Microbial community analysis At the end of the experiment, the electrodes were gently rinsed in sterile water, and the anodic and cathodic biofilms were scraped off the electrodes using a sterile spatula before being resuspended in 10 mM TriseHCl pH 8.0. A 50 mL aliquot of the cathodic biofilm was fixed and refrigerated for flow cytometry analysis. The cells were centrifuged at 10 000 g for 30 s and the pellets stored at 20  C. Microbial DNA was subsequently extracted using a PowerSoil DNA Isolation kit (MO Bio Laboratories) according to the manufacturer's directions. DNA quantity and purity was evaluated using a Nanodrop-1000 Spectrophotometer (NanoDrop 1000;

Thermo Scientific, USA). To provide phylogenetic identification of bacterial operational taxonomic units (OTUs) within each biofilm, 16S rRNA amplicon sequencing was employed. DNA was amplified using the 27F (AGAGTTTGATCMTGGCTCAG, [31]) - 519R (GWATTACCGCGGCKGCTG, [32]) primer pair, which targets the V1-V3 variable regions of the 16S rRNA gene. Amplicon sequencing was subsequently performed using the Illumina MiSeq platform (Ramaciotti Centre for Genomics; Sydney, NSW, Australia) following the manufacturer's guidelines. Raw data files in FASTQ format were deposited in NCBI Sequence Read Archive (SRA) with the study accession number SRP097212 under Bioproject number PRJNA357293. Sequences were analysed using the Quantitative Insights into Microbial Ecology (QIIME) pipeline [33]. Briefly, paired-end DNA sequences were joined, de novo OTUs were defined at 97% sequence identity using UCLUST [34] and taxonomy was assigned against the Greengenes database (version 13/8/2013) using BLAST [35]. Chimeric sequences were detected using ChimeraSlayer [36] and filtered from the dataset. Sequences were then rarefied to the same depth to remove the effect of sampling effort upon analysis. 2.6. Flow cytometry The ratio of algal-to-bacterial cells in the catholyte and biofilm was assessed by flow cytometry. A 50 mL aliquot of the freshly scraped cathodic biofilm and 1 mL of catholyte was fixed with glutaraldehyde (final conc. 2%) and stored in the dark at 4  C for less than 12 h. Prior to analysis, the samples from the two replicates were stained with SYBR-Green-I (final concentration 1:10 000) and incubated for 15 min in the dark before being diluted (1:1000) in a 0.22 mm filtered phosphate-buffered saline (PBS 1) solution. Fifty microliter of diluted samples were quantified on a flow cytometer (BD Accuri C6; BD Biosciences) with a flow rate of 35 mL min1. Bacterial and algal populations were discriminated according to cell side scatter (SSC-A) and green fluorescence (FL1-A) [37]. 3. Results and discussion 3.1. Microbial fuel cell performance with different catholytes This work aims to estimate the effect of C. vulgaris on the power

Fig. 2. Current density produced by the microbial fuel cells, MFC1 (in grey) and MFC2 (in black), during the enrichment phase with an anode potential of 0.36 V vs Ag/AgCl controlled by a potentiostat. The insert shows the start-up time after inoculation. The arrow indicates the switch from ethanol medium to synthetic wastewater (4%) and the cross indicates the time at which the cyclic voltammogram (Fig. 6a) was performed. Current density was calculated from the projected anode surface area.

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Fig. 3. Power density generated by the MFCs using the following cathode media: phosphate buffer (1 M) from days 0e15, 4 day-old anodic effluent from days 15e20, and 4 day-old anodic effluent with C. vulgaris from days 20e31. The anode and the cathode were connected via a 1 kU resistor. The anolyte and the catholyte were replenished with fresh SWW4% and “fresh” anodic effluents every 3e5 days, respectively. The red arrows indicate when the power curves were performed, and the cross indicates when the cyclic voltammetry and sampling for the flow cytometry and 16S rRNA sequencing were carried out. Data are the average of the two biological replicates. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

output of MFCs, wastewater nutrient removal and on the cathodic bacterial community composition. It should bring useful knowledge into the field of MFCs and broaden the range of applications. A measurable current was produced 110 h after inoculation of the MFCs with freshwater sediment in the two replicates, as shown in the insert in Fig. 2. The current density increased after feeding of the anodic biofilm with fresh ethanol medium, and then decreased as the biofilm depleted the nutrients within the medium. After one month, the microbial fuel cells were steadily producing an average maximum current density of approximately 1.3 A m2 (projected anode surface area of 0.00112 m2, Fig. 2), which dropped to about 1 A m2 when ethanol medium was replaced with 4% synthetic wastewater (SWW4%). Fig. 3 shows the power density generated by the MFCs after two months of anodic biofilm enrichment. The power density delivered by the system in a 1 kU resistor-configuration with phosphate buffer at the cathode was about 25 mW m2 (Fig. 3). The performance of the microbial fuel cells declined when the catholyte was changed from phosphate buffer to anodic effluent. The addition of C. vulgaris to the anodic effluent boosted the power density delivered by the system, especially under illumination (Fig. 3). In the presence of C. vulgaris, power production followed a diurnal pattern with an increase during the day and a decrease at night. The diurnal cycling of the power output was closely related to the dissolved oxygen concentration in the catholyte, as shown on Fig. 4. Under illumination, the alga photosynthesized and released oxygen into the catholyte. The oxygen produced was then reduced to water at the cathode surface and contributed to an increase in power density. During the 12 h of darkness, C. vulgaris respired the oxygen, resulting in a decrease in dissolved oxygen concentration and therefore, a decrease in power density (Fig. 4). It is likely that the bacterial community associated with C. vulgaris also contributed to the decrease in oxygen concentration by respiration, independently of the light regime. Prior studies have reported the influence of light and dark cycles on photosynthetic MFCs with C. vulgaris at the cathode [23,24,38,39]. These studies showed low

to no power production during the dark phase for C. vulgaris grown in optimal medium (Bold's basal medium or BG-11). The response in oxygen concentration was slightly slower than the power density response (Fig. 4). The cell density measurement, performed by flow cytometry at the end of the experiment, revealed that amongst all C. vulgaris cells present in the MFC (combining both biofilm and the catholyte; excluding cells attached to other surfaces in the cathode chamber), 90% were attached to the electrode surface, while 10% were in suspension in the catholyte. Therefore, the majority of C. vulgaris cells were present in close proximity to the cathode surface, as illustrated by the green cathodic biofilm in Fig. 1b. The dissolved oxygen concentration was measured in the catholyte, while the power density related directly to the processes happening at the cathode surface. The difference in oxygen concentration and power density responses could result from an oxygen gradient, with more oxygen being produced in the vicinity of the electrode surface before diffusing toward the oxygen probe. The maximum dissolved oxygen concentration reached during illumination period decreased over the course of the experiment from 700 to 500 mmol L1, also resulting in a lower power density (Fig. 4). The decrease in oxygen concentration was probably due to a combination of two factors: the consumption of oxygen by C. vulgaris associated bacterial community and lower photosynthetic rate due to light limitation as the algal biofilm grew thicker. Fig. 5 shows the maximum quantum yield of C. vulgaris cultures in 4 day-old anodic effluents, as an indicator of photosystem II stress. Although the algal cells on the cathode surface had limited access to light, they remained healthy. An average maximum quantum yield of 0.63 ± 0.07 was recorded over the 5 days in anodic effluents (Fig. 5). This value was in the range of the maximum quantum yields reported in the literature for C. vulgaris growing at 24  C under 200 mmol m2 s1 [40e42], demonstrating that C. vulgaris was not under significant stress while attached to the cathode surface. Before comparing the power densities delivered by the system

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Fig. 4. Relationship between the dissolved oxygen concentration (LHS axis; grey line) in the catholyte and the power density (RHS axis; black line) generated by the microbial fuel cell between day 23 and 28 (see Fig. 3). The periods of light and darkness are indicated by the white and grey bars, respectively. The data shown are from MFC1, representative of the two replicates. The power density of MFC2 for the same days is given in supplementary material for comparison.

Fig. 5. Maximum quantum yield of PSII (Fv/Fm) of the catholyte after inoculation of the cathode chamber with C. vulgaris in 4 day-old anodic effluents. Mean ± SEM is shown (n ¼ 2).

with different catholytes, it is important to make sure that the performance of the anodic biofilm did not change over time. To do so, cyclic voltammetry was performed before and after treatment with the different catholytes. Cyclic voltammetry provides information on the efficiency of a biofilm at transferring electrons to an electrode and its stability over time. Fig. 6a shows the cyclic voltammograms performed in turnover conditions before and after treatment. Both voltammograms were similar and displayed a sigmoid catalytic wave typical of substrate oxidation by anodic biofilms. The midpoint potential (EKA) of the system, which is the potential where half the maximum current is produced, decreased from 0.37 V to 0.47 V vs Ag/AgCl over the course of the experiment (Fig. 6a). The three month-old anodic biofilm has not been impaired by the switch from a potentiostat to a 1 kU resistor

configuration, and was actually performing better, as it transferred electrons to the anode at an even lower potential (Fig. 6a). Indeed, the anodic biofilm was selected to transfer electrons to the electrode at 0.36 V vs Ag/AgCl, but it could function at an even lower potential as, at the end of the experiment, the open-circuit potential of the anode was 0.53 V vs Ag/AgCl. The midpoint potentials observed were consistent with measured EKA values for Geobacter sp. [43], which is further supported by the 16S rRNA sequencing results for the anodic biofilm, where Geobacter sp. accounted for more than 50% of the most dominant bacterial orders (Fig. 7a). However, the catalytic current (Icat) decreased by about 0.5 A.m2 after treatment. The catalytic current is typical of the microbial oxidation rate; it provides data on the electron transfer efficiency of the biofilm. The biofilm was less efficient at oxidizing the synthetic wastewater (SWW4%) after treatment with the different catholytes medium, probably due to a slight change in the biofilm community structure over time, as previously observed in similar conditions [29]. Although, the biofilm was slower at sourcing the electrons from the catabolism of the complex SWW4%, it was still as efficient at transferring electrons to the anode (according to the EKA). In conclusion, the cyclic voltammograms show that the conductivity of the anodic biofilm was the same before and after treatment. Therefore, the differences in power output observed with the different catholytes were not due to changes in electron transfer rate of the anode, but rather in cathodic reaction efficiency. Turnover cyclic voltammetry of the biocathode showed an onset of the catalytic current at 0.1 V vs Ag/AgCl, similar to that obtain with the bare electrode, implying that the biocathode did not catalyse the reduction of oxygen at the cathode surface (Fig. 6b). Therefore, the higher power density observed in the presence of C. vulgaris was mostly due to a higher dissolved oxygen concentration under illumination, enhancing the abiotic reduction of oxygen at the cathode surface. The presence of C. vulgaris at the cathode boosted the maximal

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Fig. 6. Cyclic voltammograms in turnover conditions of (a) the anodic biofilm before and after treatment with the different catholytes in 1 kU-configuration; (b) cathodic biofilm enriched with C. vulgaris (dissolved oxygen concentration of 500 mmol L1) compare to a bare electrode of the same surface area. The catalytic current (Icat) corresponds to the maximal current at the plateau after the catalytic wave. The midpoint potential (EKA) is the potential where half the maximum current is produced. Voltammograms of MFC1, representative of the two replicates.

Fig. 7. Anodic (a) and cathodic (b) biofilm communities of the two MFC replicates (MFC1, MFC2) based on 16S sequencing. Communities were sampled at the end of the experiment after 2 weeks operation with C. vulgaris at the cathode. The orders of the most abundant bacteria in the communities are given in the legend.

power and lowered the internal resistance of the MFCs (Table 1), but the differences in performance of the MFC with different catholytes were not statistically significant (t.tests, P > 0.5). Surprisingly, the microbial fuel cells performed as well with phosphate buffer as with anodic effluent enriched with C. vulgaris. However, the performance was impaired when the cathode chamber was filled with anodic effluent only. During daytime, the oxygen concentration in the catholyte enriched with C. vulgaris (750 mmol L1) was three times higher than the maximum dissolved oxygen concentration obtained by

Table 1 Maximum power densities (Pmax) and internal resistance (Rint) with the different catholytes. Pmax is shown in mW m2 and mW m3, as per anode surface area of 0.00112 m2 and volume of 0.0001 m3 respectively. Means ± SE are shown (n ¼ 2). Catholytes

Pmax (mW m2)

Pmax (mW m3)

Rint (U)

Phosphate buffer þ air Anodic effluents þ air Anodic effluent þ C. vulgaris

31.6 ± 9.6 15.6 ± 6.9 34.2 ± 10.0

354.3 ± 107.8 174.7 ± 77.2 383.6 ± 112.2

802 ± 11 1018 ± 402 673 ± 113

bubbling air in the absence of the alga (250 mmol L1). The diffusion of oxygen to the cathode surface was faster as the oxygen concentration increased, therefore enhancing the cathodic reaction and lowering the internal resistance of the system. However, the MFC with the catholyte enriched with C. vulgaris did not perform significantly better than the MFC with phosphate buffer (t-tests, P > 0.05), suggesting that proton diffusion from the anode to the cathode was the rate-limiting step. Phosphate buffer was used to prevent the formation of a low pH inside the anodic biofilm, potentially leading to bacterial inhibition, and as a result, impairing the current generation [7]. Buffering the medium is especially relevant at high COD concentration (>1000 mg L1), as it encourages the growth of non-electrogenic microorganisms likely to reduce the pH while producing shortchain fatty acids under anaerobic condition, as shown by
lez del Campo et al. [39]. The pH of the phosphate buffer used Gonza at the cathode was 7.4, while the starting pH of the anodic effluent (4 day-old) was 6.7, before reaching 7.6 and 8.8 after 4 days incubation in the cathode chamber in absence and presence of C. vulgaris, respectively. Due to the high concentration of phosphate

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buffer, the pH of the catholyte enriched with microalgae did not fluctuate with the diurnal cycle (data not shown). The production of oxygen by photosynthesis increased the pH to 8.8, hence decreasing the concentration of one cathode reactant (Hþ), while increasing the other (O2). Protons had to diffuse all the way from the anode chamber, limiting the reaction. As previously mentioned, proton transfer to the cathode is often a limiting factor, usually addressed by increasing the CEM surface, the ionic strength of the catholyte, the buffer concentration, or directly pumping anolyte into the cathode chamber, as was performed here [5,13]. When operated with phosphate buffer, the dissolved oxygen concentration was the factor limiting the power output of the MFC, while the diffusion of protons from the anode was the limiting factor in the presence of C. vulgaris. The poor oxygen reducing activity of the electrode used, graphite [4], could also limit the power output, as C. vulgaris did not catalyse the reduction of oxygen (Fig. 6b). The limitation of the power output by the dissolved oxygen was even more obvious when the catholyte was solely made of anodic effluent. We observed that the dissolved oxygen concentration in the catholyte sparged with air dropped from 250 mmol L1 to 237 mmol L1 after 24 h. The high organic load of the effluent likely promoted the growth of aerobic heterotrophic microbes, competing with the cathode for oxygen, as observed by Freguia et al. [13] in a similar configuration. The lower dissolved oxygen content, coupled with biofouling of the cathode surface, could partially explain the lower power density and higher internal resistance observed with anodic effluent lacking C. vulgaris. The maximal power density of 34.2 ± 10.0 mW m2 obtained using C. vulgaris at the cathode is in the same order of magnitude as values previously reported in the literature. Gonz
alez del Campo et al. [39] recorded 23.97 mW m2, Wu et al. [24] 24.4 mW m2, Hou et al. [25] 19 mW m2, and Gouveia et al. [26] achieved a power density of 62.7 mW m2 at a light intensity of 96 mmol photon m2 s1 or 10 mW m2 at 26 mmol photon m2 s1. All these studies involved the growth of C. vulgaris in optimized culture medium (e.g. BG11 medium, Bold's basal medium with/without PBS, sparged or not with CO2). This study demonstrates that the use of anodic effluent as a growth medium for C. vulgaris does not hinder the maximal power output of the system.

Thauera (representing 63% of the Rhodocyclales order) was identified within the cathodic biofilm. Thauera sp. have been shown to contribute to the biological removal of nitrogen in wastewater treatment [45,46]. Xie et al. [47] identified a correlation between the increase in proportion of Thauera sp. and the nitrogen removal efficiency in an MFC-AnaerobiceAnoxiceOxic (AA/O) bioreactor. They also discovered that Rheinheimera sp. were significantly enriched on the cathode surface of their MFC AA/O reactor and suspected it to contribute to nitrogen removal [47]. Rheinheimera sp. were also identified here in the MFC1 cathodic community, and are members of the second most dominant order observed in this sample, namely the Alteromonadales. Several species of Rheinheimera are known to be capable of reducing nitrate [48,49] and occur widely in freshwater ponds [49] and bioreactors treating wastewater [47,50]. Although Thauera sp. and Rheinheimera sp. are not common C. vulgaris-associated genera [51,52], these groups were dominant in the cathodic environment of our system. Rhizobiales also constituted 11% of the bacterial orders present within the cathodic communities, with three families identified: Brucellaceae, Phyllobacteriaceae, and Rhizobiaceae. Many species of Rhizobiaceae are diazotrophs that fix atmospheric nitrogen gas into a more usable form such as ammonia [53]. Rhizobiaceae has been identified as one of the main bacterial families associated with Chlorella pyrenoidosa cultures [54]. It was also the most abundant family (represented by the Rhizobium genus) recovered from the inside surface of a prototype OMEGA (Offshore Membrane Enclosures for Growing Algae) photobioreactor treating municipal wastewater primarily inoculated with the green algae Scenedesmus sp. and Desmodesmus sp. [50]. Our cathodic biofilms contained putative diazotrophs (nitrogen-fixing bacteria), denitrifying and nitrate reducing bacteria, all likely to influence the nitrogen cycle at the cathode. Other dominant bacterial orders observed here, including the Flavobacteriales (Fluviicola sp) and Sphingomonadales, are groups typically associated with phytoplankton [55]. The Sphingomonadales are known to be associated with Chlorella sp. They have been repeatedly isolated from non-axenic Chlorella cultures [51,52,56e58], but their specific interaction mechanism with Chlorella sp. is still unknown. 3.3. Chemical oxygen demand and ammonium removal

3.2. Anodic and cathodic microbial communities Fig. 7 shows the formation of discrete bacterial assemblages within the anodic and cathodic biofilms. The anodic biofilm was largely dominated by Geobacter sp. as expected from the electrochemical analyses. The Bacteroidales and Clostridiales enriched at the anode in synthetic wastewater persisted at the cathode (Fig. 7). Flow cytometry performed at the end of the experiment demonstrated that the cathodic biofilm was comprised of 22 ± 7% of algal cells and 78 ± 7% of bacterial cells (ratio of 1:3.5), while in contrast the catholyte contained 2.3 ± 0.5% of algae and 97.7 ± 0.5% of bacteria in suspension (ratio of 1:42). Bacteria dominated both the cathode surface and the catholyte; so it is therefore important to identify the composition of the bacterial assemblage occurring in the presence of C. vulgaris and speculate about their respective roles. The culture of C. vulgaris used to inoculate the cathode chamber was not axenic, therefore some of the bacteria present are likely to be part of C. vulgaris associated bacterial community. Members of the Pseudomonadales dominated the cathodic biofilm (Fig. 7), with an operational taxonomic unit (OUT) matching Pseudomonas stutzeri accounting for 50% of these. Pseudomonas stutzeri is a denitrifying bacterium commonly found in soils [44], which possesses the cellular machinery to convert nitrates into free atmospheric nitrogen (N2). Another genus of denitrifying bacteria,

The chemical oxygen demand (COD) measures the concentration of organic compounds in wastewater, and is an important indicator of treated water quality. With a retention time of 4 days, the anodic biofilm depleted about 34% (0.98 ± 0.28 g L1) of the chemical oxygen demand in the original medium (initial COD of 2.9 g L1) and the cathodic biofilm removed an extra 10e15% (0.3e0.4 g L1) in the absence or presence of C. vulgaris (Fig. 8a). With a total retention time of 8 days, the rate of COD removal in the presence of the microalga was 0.19 g L1 d1 and the coulombic efficiency (CE) of the system was 13.6 ± 1.2%. This CE did not vary significantly in presence or absence of the algae, meaning that the extra COD removed at the cathode was not used in electricity generation but rather in biomass production. The stable CE also demonstrates that the anodic biofilm maintained its efficiency over time, as previously shown by cyclic voltammetry. The differences in power output observed with the different catholytes were mostly due to variations in cathodic reaction efficiency. Chlorella vulgaris is a photolithoautotroph that derives energy from light and carbon from the fixation of (inorganic atmosphericdissolved) carbon dioxide, implying that it will not directly contribute to the removal of dissolved organic carbon. However, by increasing the concentration of dissolved oxygen during daytime, C. vulgaris is likely to boost bacterial respiration, therefore

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1  Fig. 8. Measure of (a) chemical oxygen demand (COD) and (b) ammonium and nitrate (NHþ 4 and NO3 in mg L ) removal by the anode and the different catholytes. Positive and negative concentrations show production and uptake, respectively. The measurements were taken after a 4 day-incubation of the anodic biofilm in SWW4% (“Anode”), follow by another 4 day-incubation of the cathodic biofilm in anodic effluents, in presence or absence of the microalgae C. vulgaris (“Cathode without C. vulgaris”, “Cathode with C. vulgaris”). Mean ± SEM, n ¼ 2.

encouraging consumption of organic matter and lowering the COD. A wide range of COD removal efficiencies have been reported in the literature for C. vulgaris. It has been shown to vary from 20% when treating kitchen waste anaerobically digested effluent for 19 days (removal rate of 0.02 g L1 d1) [59], to 75.5% in dairy manures with a 5 days hydraulic retention time (HRT) (removal rate of 3.6 g L1 d1) [19], and 38% in textile wastewater with 10 days HRT (removal rate of 0.03 g L1 d1) [20]. None of these studies analysed the composition of microbial communities associated with C. vulgaris, even though it could greatly affect the amount of COD being removed. Factors influencing the COD removal include the design of the reactor, the microbial consortia, the growth rate of C. vulgaris (for dissolved oxygen production), the composition of the wastewater, the temperature of operation, and the retention time [18]. The overall 49% COD removal efficiency observed in this study could also be increased by effective stirring in both chambers. After 4 days of incubation of the anodic biofilm in SWW4%, the concentration of dissolved ammonium almost doubled. The production of 22 ± 1 mg L1 ammonium was also observed after the 4 day-incubation of the cathodic biofilm without C. vulgaris in anodic effluent (Fig. 8b). The production of ammonium was probably due in part to the presence of diazotrophs, shown to occur in the system, but also to derive from urea degradation, as it was the principal source of nitrogen in the synthetic wastewater. The anode was inoculated with freshwater sediments likely to contain microorganisms with urease activity, as this is very common in soil [60]. Urease is an enzyme that catalyses conversion of urea to ammonia, ammonium and bicarbonate. Less ammonium was produced in the catholyte lacking C. vulgaris than in the anode chamber, because the anodic community had already assimilated some of the urea. The presence of C. vulgaris at the cathode improved the removal of ammonium, with a reduction of 24 ± 8 mg L1 in ammonium concentration. Taking into account the approximately 20 mg L1 of presumed ammonium “production” due to urea degradation, C. vulgaris could actually be consuming more than 40 mg L1 of ammonium, corresponding to a removal rate of 5 mg L1 d1 with an 8 days retention period. The ammonium removal rates reported in the literature for C. vulgaris growing in wastewater vary from rates as high as 200 mg L1 d1 [19,21] to rates as low as

0.65 mg L1 d1 [20], likely to depend on the growth rate of C. vulgaris in the type of wastewater tested. The observed reduction in ammonium concentration could be the result of direct assimilation by C. vulgaris, and/or by the bacterial community associated with it. The microbial profile (Fig. 7b) revealed the presence of denitrifying bacteria likely to contribute to the conversion of ammonium into nitrogen gas. However, since ammonium was the principal source of nitrogen in the system, the absence of ammonium-oxidizing (e.g. Nitrosomonas genus) and nitrite-oxidizing bacteria (e.g. Nitrobacter genus) within the cathodic biofilm was surprising. There is still a possibility that these genera were present in the catholyte but not at the electrode surface, as only the biofilm community was characterized in this study. Ammonia-oxidizing archaea could also have contributed to the uptake of ammonium [61], but the bacteria 16S rRNA primers employed here are not optimized for archaea identification. The nitrate concentration remained relatively constant during the course of the experiment, no significant difference in nitrate “production” between the different catholytes was observed. When grown with ammonium nitrate as the nitrogen source, C. vulgaris preferentially assimilates the more reduced form of nitrogen (ammonium in this case) [62]. The bacterial community profiling revealed the presence of denitrifying bacteria (P. stutzeri and Thauera sp.), but they seem to have had a negligible effect on the nitrate concentration in this system. The nitrogen-fixing bacteria present at the cathode (Rhizobiaceae) catalyse the reduction of N2 into NH3 using the nitrogenase enzyme. Members of this group have regularly been shown to occur in association with microalgae [50,54]. Even though N2 fixation is  unlikely to occur under NHþ 4 and NO3 replete conditions, the abundance of nitrogen-fixing bacteria should be limited in a system aiming to reduce the nitrogen content of wastewater. It is thus important to report the bacterial community associated with microalgae in systems shown to treat wastewater efficiently, so strategies to reproduce or engineer cathodic bacterial communities  to increase COD and N removal (as NHþ 4 and NO3 ) can be developed. To efficiently treat wastewater and produce energy in a configuration where the anode effluent is used as cathode influent in photo-cathode MFCs, several parameters have yet to be

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optimized: (i) the retention time has to be optimal to maximize COD and N removal and limit the formation of a pH gradient between the anode and the cathode; (ii) a shorter dark phase could be considered to maintain a high power density for longer; (iii) CO2 may be added to the catholyte to counter Hþ diffusion limitation, as suggested by Fornero et al. [63], while optimizing Chlorella growth and therefore, electricity production; (iv) the bacterial community should be engineered to promote algae growth and nutrients uptake at the cathode. Reporting simultaneously the microbial community composition, the electrochemical data and nutrient removal results, is essential to improve the process and increase reproducibility between studies. 4. Conclusions In this study, microbial fuel cells were operated using the anode effluent as cathode influent to counter pH gradient limitation and polish COD removal. Chlorella vulgaris was grown in anodic effluent at the cathode to provide oxygen via photosynthesis and increase the power output. This system resulted in several advantages over conventional MFC designs. The presence of C. vulgaris doubled the maximum power density compared to the power generated without the alga, and largely contributed to the removal of ammonium. The power output closely followed the production of oxygen and cycled diurnally. The bacterial community associated with C. vulgaris was also investigated, revealing the presence of bacteria involved in nitrogen cycling and highlighting the importance of characterizing these consortia to enable community engineering strategies. Overall, the system described here reduces the use of expensive cathode catalysts and catholytes, enables the production of electricity and algal biomass, while contributing to the removal of wastewater pollutants. Even though further optimisation is needed, similar systems could be implemented in next generation wastewater treatment plants driven by biological recovery technologies.

[9] [10] [11] [12] [13] [14] [15] [16] [17] [18]

[19] [20] [21] [22] [23] [24] [25] [26] [27] [28] [29] [30] [31]

[32] [33]

[34] [35]

Acknowledgements The project was funded by the Climate Change Cluster of the University of Technology Sydney. The authors would like to thank Dr Jean-Baptiste Raina for his help with the flow cytometer, Dr Milan Szabo for his guidance with the pulse-amplitude modulated fluorometer, and Dr Md Abu Hasan Johir for his assistance with the ion chromatograph. The authors are also thankful for Dr Helen Price's help with the Gallery™ Plus Automated Photometric Analyzer. Appendix A. Supplementary data Supplementary data related to this article can be found at http:// dx.doi.org/10.1016/j.jpowsour.2017.03.097.

[36]

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