Enrichment of electrochemically active bacteria using microbial fuel ...

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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