Enrichment of electrochemically active bacteria using microbial fuel cell and potentiostat

Enrichment of electrochemically active bacteria using microbial fuel cell and potentiostat Tim Niklas Enke ETH Zurich timenke@hotmail.com tim.enke@usys.ethz.ch Microbial Diversity 2015

Introduction Microbial fuel cells (MFC) can be applied to harness the power released by metabolically active bacteria as electrical energy (Figure 1). In addition to the energy generation capabilities of MFC, they have been used to generate hydrogen gas and to clean, desalinate or detoxify wastewater [1,2]. Among the bacteria found to be electrochemically active are Geobacter sulfurreducens, Shewanella putrefaciens and Aeromonas hydrophila [2,3,4]. Figure 1: Scheme of a microbial fuel cell. A MFC consists of an anaerobic anode chamber with rich organic matter, such as sludge from wastewater treatment plants or sediment. The anode (1) serves as an electron acceptor in an electron acceptor limited environment and is wired externally (2) over a resistor (3) to a cathode (5). Electrons travel over the circuit and create a current, while protons can pass the proton exchange membrane (4) to reach the oxic cathode chamber. At the cathode, the protons, electrons and oxygen react to form water. In the cathode chamber, a catalyst can facilitate the reaction and thus the movement of electrons. Figure from [https://illumin.usc.edu/assets/media/175/mfcfig2p1.jpg, 08/18/2015]. Even more remarkably, in the deep sea, microbes can power measurement devices that deploy an anode in the anoxic sediments and position a cathode in the oxygen richer water column above, thus exploiting the MFC principle [5]. In a different application, MFC can be used to enrich for bacteria that are capable of extracellular electron transfer (EET) and form a biofilm on the electrode. In this setup, the MFC anode serves as an electron acceptor in a rich

organic, anaerob environment that is limited for electron accepting species, providing a niche and thus selecting for EET capable bacteria [6]. Contrary to an MFC where electrochemically active bacteria are enriched due to their capability to donate electrons to an anode, a potentiostat sets a constant potential between a working and a reference electrode by adjusting the current. Here, the enrichments selects for bacteria that are capable of using electrons to harvest energy. Furthermore, potentiostats can be used for cyclic voltammetry, where a potential is cycled and the resulting current is recorded to investigate redox chemical processes at the working electrode. This mini project aims at probing the potential of microbial fuel cells and potentiostat to in situ and in vitro enrich for electrochemically active microbial consortia. Results Graphite electrodes were incubated in a microbial fuel cell (see Figure 5, also Figure 1), in vitro in a core from Trunk river (Figure 4) and in situ at Trunk river (Table 3). The electrodes and controls from the MFC, the core (no controls) and in situ site at Trunk river (no controls) were imaged with a stereoscope to look for biofilm formation and for some electrodes cyclic voltammetry was performed to investigate the redox activities on the electrode (Table 1). Parts of the electrodes were fixed and prepared for scanning electron microscopy to further investigate biofilm composition (Table 2). Microbial fuel cell The potential between anode and cathode was measured for eight days (Figure 2). In the microbial fuel cell, an increase in potential can be observed, plateauing after 5 days. The anode used to enrich for bacteria capable of EET shows a different biofilm than the control that was deposited in the anode chamber of the MFC but not wired to a cathode, thus it just provided a graphite surface and no electron sink (Table 1, b and c). Scanning electron microscopy showed that the biofilm on the anode consists of both larger single cell eukaryotes as well as small round bacteria in a dense biofilm with extracellular matrix (Table 2, b).

The MFC anode was re-inoculated into a fresh MFC with glucose / galactose media and media composition, OD and potential were monitored over time (Figure 3). While OD increase to 0.3, the potential did not show any increase. After three days, no more OD increase was observed and the anode was harvested. The biofilm on the anode from the secondary enrichment is different from the biofilm that grew on the anode from the first enrichment (Table 1 d). Consistent with the decreasing potential in the secondary enrichment, the anode did not show any redox activities in cyclic voltammetry. Figure 2 Microbial fuel cell and core potential between the anode and the cathode. a) b) c) d) Figure 3 Secondary enrichment: the anode from the MFC was re-inoculated into a fresh MFC setup and monitored. a) OD over time b) potential between the anode and the cathode over time c) consumption of glucose and galactose in MFC medium d) production of galactose and glucose break down products, c) and d) monitored by HPLC.

Trunk river in vitro core The core reached an equilibrium potential after 40 hours and showed no increase in potential (Figure 2). The observed biofilm on both the cathode and the anode appeared different, but showed no redox activity in cyclic voltammetry measurements (Table 1 g and h), consistent to the equilibrating and not increasing potential measurement. Figure 4 Oxygen and hydrogen sulfide profiles for the first 4.5 cm of the sediment of core from trunk river, determined with microelectrodes. The core contains an anode in the sediment (ca. 12 cm deep, presumably in the anaerobe region) and the cathode at the air water interface. Trunk river in situ electrode enrichments Electrodes were harvested from the in situ site at trunk river after 12 days, although the anodes were lost due to cable corrosion in 3 out of 4 cases. A different biofilm on cathode and anode can be observed (Table 1 e,f). SEM of the electrodes show many large cells on the cathode and a dense bacterial biofilm on the anode (Table 2 c, d). Two of the cathodes were re inoculated into anaerobic bottles with Fe2+ containing medium to check if the enriched bacteria can oxidize and accept electrons from iron, both under light and dark conditions. The incubations appeared orange as a sign of iron oxidation and the electrodes were harvested after 8 days and investigated by microscopy and cyclic voltammetry (Table 1, k and l). Both electrons show a very different biofilm and redox activity in the cyclic voltammetry.

One cathode from trunk river was used to inoculate a potentiostat and harvested after 8 days of constant potential. Compared to the reference electrode, the region of the cathode that was submerged in the potentiostat media showed a clear biofilm (Table 1, i and j). In addition, cyclic voltammetry revealed redox activity on the potentiostat electrode. Table 1 Stereoscope images and cyclic voltammetry profiles (if available) of electrodes from different enrichments. Source electrode Image Cyclic voltammetry a) control graphite control b) MFC first enrichment anode c) MFC first enrichment control

d) MFC second enrichment anode e) Trunk river cathode f) Trunk river anode g) Core Trunk River cathode

h) Core Trunk River anode i) Potentiostat electrode Working electrode j) Potentiostat electrode Counter electrode (ctrl) k) Fe2+ light cathode

l) Fe2+ dark cathode Table 2 Scanning Electron Microscopy images of electrodes from different enrichments Source electrode SEM image a) control graphite control

b) MFC first enrichme nt anode

c) Trunk River cathode

d) Core Trunk River anode Discussion The different biofilms on the electrodes show that different inoculum sources as well as the different enrichment procedures lead to the formation of distinctable biofilms. Stereomicroscopy yields a variety of different biofilm types that grow on the graphite electrodes from different sources and cyclic voltammetry confirmed redox activity of some of the biofilms. Scanning electron microscopy revealed both bacterial biofilms as well as associated diatoms and other larger single cell organisms, specifically at the cathodes from trunk river. The potentiostat caused a biofilm to develop on the working electrode that showed peaks of redox activity in cyclic voltammetry. To conclude, both the MFC and the potentiostat setup allow to enrich for and study electrochemically active bacteria that form biofilms on the electrodes. Apart from the here applied methods used to investigate the electrodes, stereomicroscopy, scanning electron microscopy and cyclic voltammetry, other methods can give complementary insight: FISH can reveal the phylum composition of the consortia as well as the spatial organization within the biofilm. Plating on indicator plates like MnO2 plates that clear upon electron transfer to the MnO2 can help to isolate and further characterize bacteria capable of extracellular electron transfer.

Caveats in the experimental setup were corrosion of in situ electrode cables in trunk river that were not insulated. Corrosion can decrease the conductivity of the cable and in this case even caused the breaking of the wire and loss of the anodes. Furthermore, controls that were not wired to a circuit to investigate biofilm formation on graphite in the absence of electron transport were only included in the MFC and not in the in situ samples. Including controls and insulating the cables that connect the electrodes can lead to more conclusive insights in the biofilm formation at mfc electrodes. For the secondary MFC enrichment, the membrane could not be fully recovered and was covered by a white film. Even harsher cleaning conditions did not result in a clean membrane. If the membrane was not permeable for protons in the second set up, the declining potential in the second enrichment can be explained. In parallel to the presented MFC, three do it yourself MFC with different sediments as inoculum were set up to compare differences in biofilm formation at the anode (see for example http://www.engr.psu.edu/ce/enve/logan/bioenergy/mfc_make_cell.htm). These MFC used an agar saltbridge instead of a membrane, but none of them created a change in potential, which can be because of the high internal resistance of the saltbridge or oxygen leakage into the anaerobic anode chamber. Still, the anodes graphite electrodes showed biofilm formation even for the self-made MFC (data not shown), although a conclusion whether these are electrochemically active bacteria is not possible without an increase in potential.

Methods and Protocols Table 3 Inoculum sources for MFC and core Inoculum source Sippewissett Salt Marsh (SW) Trunk River (TR) Description intertidal salt marsh, photosynthetic microbial mats, multicellular Magnetotactic Bacteria (MMBs) Trunk River freshwater / brackish basin overlying sediments with seawater intrusion and an active sulfur cycle MFC set up Proton Exchange Membrane MFC Core, in situ electrodes Microbial Fuel Cell setup Figure 5 Microbial Fuel Cell setup, secondary enrichment. Left: anaerobic anode chamber with MFC media, gas outlet and bubbled with nitrogen. Proton exchange membrane between the two chambers. Right: cathode chamber with 50 mm Potassium ferrycyanide in 1:1 SW and FW base as catalyst, bubbled with air. See also Figure 1. Electrodes are 2.5-3 cm graphite with a hole drilled with syringe needle. Wire used throughout was copper cable. The cable was insulated with rubber coating (Performix Plasti Dip) to prevent corrosion.

The aerobe cathode chamber contained 50 mm of the catalyst potassium ferricyanide (K3[Fe(CN)6 to facilitate electron acceptance by oxygen (2H+ + 2e- + O2 -> H2O). The cathode is wired over a 220 Ohm resistor to the anaerobe anode chamber. The secondary enrichment MFC was set up as stated above. Inoculum was the anode from the first enrichment. Microbial Fuel Cell Media for second enrichment Ingredient and stock conc Final conc. 500 ml SW base 1 x 10 ml 100 x FW base 1 x MOPS, ph 7.2, 1M 20 mm Galactose 1M 10 mm Glucose 1M 10 mm NH4Cl 100 x 10 mm H2S 1M 1 mm K2HPO4 100 mm 1mM Trace Elements and Vitamins 1x Proton Exchange Membrane preparation (protocol provided by Lina Bird) a. To clean membranes, place all dirty membranes in 70% ethanol solution for 30 minutes. i. Ethanol cleans off grease & graphite fibers from membranes. b. Wipe off grease from membranes using ethanol and kimwipes. After removing grease, immediately place each membrane in a beaker of DDI water. i. Membranes should be in solution at all times to prevent drying and cracking. c. Rinse with fresh DDI water. d. Boil membranes on low (~80C) in ddh2o for 30 minutes. Rinse. e. Boil membranes on low (~80C) in 3% H2O2 for 1 hour. Membranes will often float above fluid line weigh down the membranes with a glass apparatus to keep them submerged. i. H2O2 cleans the membrane. f. Rinse thoroughly with DDI water. g. Boil membranes on low (~80C) in 0.5 M H2SO4 for 1 hour. See notes in Step E. i. H2SO4 re-protonates membranes & provides additional cleaning. h. Rinse thoroughly in DDI water. i. Store in DDI water in Clean Membranes container. j. If pretreating new membranes, cut membranes out to dimensions of 5 x 5 cm. Soak in 0.5% HCl for 2 3 hours. Rinse with DDI water. Follow steps D H. Store in DDI water in New Membranes container.

Iron media (Fe 2+ ) Ingredient and stock conc Final conc. 10 ml 100 x FW base 1 x MOPS, ph 7.2, 1M 20 mm Acetate 1M 1 mm Bicarbonate 1M 25 mm NaNO3 10 mm Fe 2+ 5 mm NH4Cl 100 x 10 mm NaSO4 1M 1 mm K2HPO4 100 mm 1mM Trace Elements and Vitamins 1x Potentiostat media Ingredient and stock conc Final conc. 500 ml SW base 1 x 10 ml 100 x FW base 1 x NH4Cl 100 x 10 mm Bicarbonate 1M 25 mm NaSO4 1M 1 mm K2HPO4 100 mm 1mM Trace Elements and Vitamins 1x Fixation for SEM Electrodes were submerged in 4 % PFA and incubated 4h at 4 C. After fixation, sampled were washed 3 times in 1x PBS and dehydrated by each 20 minutes at room temperature in 25%, 50%, 75%, 95% and 100% ethanol. Samples were further dried by critical point drying and spotter coated with platinum in the MBL central microscope facility.

Acknowledgements I want to thank the Bernard Davis Endowed Scholarship Fund and ETH Zurich for the financial support of my participation in the course. I also want to thank my supervisor Otto Cordero who encouraged my application for this course, knowing about the impact that it can and will have on every researcher s life and career. Thanks to Lina Bird for the equipment, help and discussion for the setup of the enrichments that form the basis of this mini project. Special thanks go to all the students in the course for making the intense time and experience of the Microbial Diversity course 2015 so fruitful and memorable, to all the teaching assistants who avidly worked to create a perfect working and learning atmosphere in the course, to the course assistants and the course coordinator for keeping things running and to the faculty for their advice, guidance and discussion. Lastly, both directors deserve the highest appreciation and admiration for the organization and realization of the course and the inspiration and scientific spirit they transmit on to young scientists in word and deed. References 1. Logan, B. E., Hamelers, B., Rozendal, R., Schröder, U., Keller, J., Freguia, S., Rabaey, K. (2006). Microbial fuel cells: Methodology and technology. Environmental Science and Technology, 40(17), 5181 5192. doi:10.1021/es0605016 2. Liu, H., & Logan, B. E. (2004). Electricity generation using an air-cathode single chamber microbial fuel cell in the presence and absence of a proton exchange membrane. Environmental Science & Technology, 38(14), 4040 4046. doi:doi 10.1021/Es0499344 3. Kim, B.H.; Kim, H.J.; Hyun, M.S.; Park, D.H. (1999a). "Direct electrode reaction of Fe (III) reducing bacterium, Shewanella putrefacience" (PDF). J Microbiol. Biotechnol 9: 127 131. 4. Pham, C. A.; Jung, S. J.; Phung, N. T.; Lee, J.; Chang, I. S.; Kim, B. H.; Yi, H.; Chun, J. (2003). "A novel electrochemically active and Fe(III)-reducing bacterium phylogenetically related to Aeromonas hydrophila, isolated from a microbial fuel cell".fems Microbiology Letters 223 (1): 129 134. doi:10.1016/s0378-1097(03)00354-9 5. Gong, Y., Radachowsky, S.E., Wolf, M., Nielson, M.E., Girguis, P.R., and Reimers, C.E. 2011. Benthic Microbial Fuel Cell as Direct Power Source for an Acoustic Modem and Seawater Oxygen/Temperature Sensor System. Environmental Science and Technology 45(11):5047-5053. doi:10.1021/es104383q. 6. Meng, T., Li, S., Du, Z., & Li, H. (2007). Enrichment of an Electrochemically Active Bacterial Community. Proceedings of ISES Solar World Congress 2007: Solar Energy and Human Settlement, 2434 2438