Enrichment of electrochemically active bacteria
using microbial fuel cell and potentiostat
Tim Niklas Enke
ETH Zurich [email protected] – [email protected]
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.
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.
a) b)
c) d)
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
Description MFC set up
Sippewissett
Salt Marsh (SW)
intertidal salt marsh,
photosynthetic microbial mats,
multicellular Magnetotactic
Bacteria (MMBs)
Proton Exchange
Membrane MFC
Trunk River (TR) Trunk River – freshwater / brackish
basin overlying sediments with
seawater intrusion and an active
sulfur cycle
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 (Fe2+)
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
Fe2+ 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.
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