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Electrochemical studies of outward and inward extracellular electron transfer by microorganisms from diverse environments
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Electrochemical studies of outward and inward extracellular electron transfer by microorganisms from diverse environments
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Content
ELECTROCHEMICAL STUDIES OF OUTWARD AND INWARD
EXTRACELLULAR ELECTRON TRANSFER BY MICROORGANISMS
FROM DIVERSE ENVIRONMENTS
By
Karla Abuyen
A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(MOLECULAR BIOLOGY)
August 2024
Copyright 2024 Karla Abuyen
ii
Dedication
To Jan Amend, who valued and prioritized the next generation of scientists.
iii
Acknowledgements
I would like to express my deepest gratitude to the individuals who have played
instrumental roles in my journey as a PhD student. Firstly, I am indebted to my research
advisor, Moh El-Naggar, whose unwavering support, guidance, and expertise have been
invaluable throughout this endeavor. His kindness, humility and intelligence have been a
constant source of inspiration. Throughout our interactions, he has exemplified the
qualities of empathy, generosity, and wisdom, serving as an excellent role model both for
personal and professional growth. I am honored to have had the opportunity to learn from
such esteemed scholar and mentor. More importantly, I am truly fortunate to have crossed
paths with him and deeply thankful for his friendship and positive influence.
I am immensely grateful to Jan Amend, my undergraduate research advisor. He
was a remarkable individual whose influence and guidance have left an indelible mark on
my life and career. To say the least, I won’t be here if he didn’t pave the way and provided
me (and many others) with the opportunity to explore the field of Geomicrobiology. I am
especially grateful to him for believing and trusting me in mentoring my younger peers,
it pushed me beyond my limits and challenged me to strive for excellence. I am eternally
grateful for the time I spent in his lab and will cherish the memories and lessons learned.
Although he is no longer with us, his legacy lives on in the countless lives he’s touched
and the knowledge he imparted.
iv
Furthermore, I extend my appreciation to the faculty members and peers who have
contributed to my academic and personal development through engaging discussions,
constructive criticism, and camaraderie. I am thankful to my mentors, Annie Rowe, Ileana
Perez-Rodriguez, Roman Barco, and Pratixa Savalia, their unwavering support, wisdom,
and encouragement empowered me to develop my skills as a scientist and mentor during
my time as an undergraduate in Jans lab. Thank you for teaching me the basics, you have
contributed so much to my success. To my fellow lab members, our lunches together and
walks to Dulce will hold a special place in my heart, I was always left energized interacting
with all of you. Thank you for your support as I was preparing for my dissertation, I am
lucky to have been surrounded by great individuals. I would also like to thank the
members of my qualification and dissertation committee, Steve Finkel, Vadim Cherezov,
James Boedicker and Doug LaRowe, thank you so much for your thoughtful feedback,
constructive criticism, and unwavering commitment to academic excellence. Your
collective expertise, diverse perspectives, and dedication to scholarly inquiry have enriched
my dissertation. My special thanks to Doug LaRowe who stepped up when called upon
last minute to serve as a member of my dissertation committee. His willingness to dedicate
his time, expertise, and insights to the evaluation of my research is deeply appreciated. I
would also like to thank Steve Finkel and the PhD Academy team. I am not the public
speaking type, but you guys gave me countless opportunities to overcome my fear. Thank
you for believing in me and the things I had to say. I am immensely grateful to the
dedicated team who organized and facilitated the events. Their tireless efforts and
v
commitment to the students’ personal growth have provided us with invaluable
opportunities to enhance our skills, expand our networks, and prepare for our future
careers.
I am grateful to my family and friends for their encouragement, love, and
understanding, which have sustained me during the challenges of graduate school. I would
like to thank the Abuyen family; their sacrifices and support have allowed me to pursue
my passion and navigate the complexities of academic life with greater confidence and
resilience. They are my greatest source of strength and inspiration, and I am honored to
have them by my side as I embark on this next chapter of my journey. To my friends
John and Z, thank you for hanging around and keeping me sane. To my childhood friends,
Julius, Kinna, Laila, and Niña, thank for your support from 11,594 km away. I am
profoundly grateful for your presence in my life, and I cherish the moments we have shared
together amidst the demands of graduate school. I would also like to thank my colleagues
at Viridian Biometals for their support during the last two years of my PhD and
throughout this dissertation process. Lastly, I want to thank Adam, my partner in crime
and best friend, who despite my occasional struggles in articulating complex research
concepts, he patiently listened to me discuss my work, providing invaluable
encouragement and understanding.
vi
Table of Contents
Dedication ...................................................................................................................ii
Acknowledgements ....................................................................................................iii
List of Figures............................................................................................................ix
List of tables .............................................................................................................xv
Abstract...................................................................................................................xvi
Chapter 1: Introduction..............................................................................................1
1.1 Microbiological perspective of biodiversity ....................................................................1
1.2 Oxidative phosphorylation ............................................................................................2
1.3 Extracellular electron transfer.......................................................................................6
1.4 Extracellular electron uptake ......................................................................................10
1.5 Microbial electrochemistry ..........................................................................................12
1.6 Microbes in anoxic environments.................................................................................16
1.7 The Human Microbiome..............................................................................................18
Chapter 2: Soluble Iron Enhances Extracellular Electron Uptake by Shewanella
oneidensis M R-1 ....................................................................................................... 22
2.1 Abstract......................................................................................................................23
2.2 Introduction ................................................................................................................23
2.3 Materials and Methods................................................................................................26
2.3.1Cell cultivation.....................................................................................................26
2.3.2Reactor setup and electrochemical measurements ................................................27
2.3.3Biofilm formation.................................................................................................28
2.3.4FeCl2 addition......................................................................................................28
2.3.5Iron chelator addition and toxicity test................................................................29
2.3.7Fluorescence microscopy ......................................................................................30
2.3.7Tetramethylbenzidine heme stain SDS-PAGE protein gel....................................30
2.4 Results and Discussion ................................................................................................31
vii
2.4.1Iron (II) chloride enhances extracellular electron uptake by S. oneidensis. ...........31
2.4.2 Fe-enhanced cathodic electron uptake in S. oneidensis is concentration dependent
and due to free Fe ions................................................................................................36
2.4.3 Fe-mediated electron uptake in S. oneidensis is linked to fumarate reduction .....41
2.4.4 Fe-mediated electron uptake is dependent on outer membrane and periplasmic
cytochromes ................................................................................................................43
2.5 Conclusions...................................................................................................................47
Chapter 3: Investigation on M icrobe M ineral interaction using electrochemical
cultivation at the former Homestake Gold mine in South Dakota .......................... 49
3.1 Abstract .......................................................................................................................50
3.2 Introduction..................................................................................................................51
3.3 Materials and Methods .................................................................................................55
3.3.1 Site description ...................................................................................................55
3.3.2 Geochemical analysis...........................................................................................55
3.3.3 Mineral and electrochemical incubation ..............................................................58
3.3.4 Electrochemical data analysis..............................................................................62
3.3.5 Sample collection and DNA extraction................................................................64
3.3.6 Microbial community analysis.............................................................................65
3.4 Results and Discussion..................................................................................................67
3.4.1 Evidence of both oxidizing and reducing microbes (1st incubation) .....................67
3.4.2 Changes in in situ microbial activity over time (2nd incubation) .........................78
3.4.3 Similar microbial community composition observed across treatments................81
3.5 Conclusion....................................................................................................................86
Chapter 4: Electrochemical investigation of Aggregatibacter actinomycetecomitans
D7S-1 ........................................................................................................................ 89
4.1 Abstract .......................................................................................................................89
4.2 Introduction..................................................................................................................90
4.3 Methods........................................................................................................................94
4.3.1 Cell Growth Conditions ......................................................................................94
4.3.2 Reactor setup and electrochemical measurements ...............................................94
4.3.3 Media exchange and spent medium analysis........................................................96
4.3.4 SEM Imaging ......................................................................................................97
viii
4.4 Results..........................................................................................................................98
4.4.1 Replacement of reactor media abolishes Aggregatibacter actinomycetecomitans
anodic current .............................................................................................................99
4.4.2 Spent media contains redox-active molecules .................................................... 101
4.4.3Flavin mononucleotide has no effect on the EET activity of A.
actinomycetecomitans................................................................................................ 101
4.4.4 Chemical and physical inhibition has no effect on anodic current production of A.
actinomycetecomitans................................................................................................ 102
4.4.4 An OMC Shewanella mutant has higher anodic current compared to A.
actinomycetecomitans D7S-1 ..................................................................................... 105
4.5 Discussion ................................................................................................................. 106
4.6 Conclusion ................................................................................................................ 108
Chapter 5: Conclusions........................................................................................... 111
Chapter 6: Bibliography ......................................................................................... 115
ix
List of Figures
1.1 Schematic representation of oxidative phosphorylation in the mitochondria.
Electrons transferred through the electron transport chain in the inner
mitochondrial membrane, generating a proton gradient that drives ATP
synthesis by ATP synthase. This process is crucial for cellular energy
production in aerobic respiration. Illustration from (Shrestha, 2003).
1.2 Schematic of redox tower showing electron donors and acceptors used by
microorganisms. Electron donors and acceptors are arranged with respect to
their standard reduction potential at pH 7. Electron donors with a more
reducing potential (more negative Eo’, standard conditions) are at the top of
the tower while the electron acceptors with a more oxidizing potential (more
positive Eo’) are localized at the bottom. Arrows indicate electron
donor/acceptor pairs that microbes prefer for survival. Figure courtesy of Moh
El-Naggar.
1.3 Schematic representation of extracellular electron transfer in dissimilatory
metal reducing bacteria Shewanella oneidensis MR-1. Electron transfer to the
extracellular space facilitated by inner membrane and periplasmic redox-active
proteins and is handed off to outer membrane multiheme cytochromes.
1.4 Extracellular electron transfer to solid surfaces is enabled by outer membrane
cytochromes directly interacting with solid surface, cytochrome bound
cofactors, and nanowire also known as membrane extensions consisting outer
membrane cytochromes or indirectly via soluble electron shuttles. Figure
adapted from (Chong et al., 2018).
1.5 Proposed mechanisms of extracellular electron uptake from a cathode. (A)
mediated electron transfer (B) mediated electron transfer via hydrogenase
generated H2 (C) Direct electron transfer through outer membrane
cytochromes.
1.6 Schematic of the bioelectrochemical reactor. The reactor consisted of a
stoppered glass bottle with a graphite working electrode (WE), platinum
counter electrode (CE), Ag/AgCl reference electrode (RE), and N2 ports.
1.7 Electrochemical techniques used in this study. Top figure illustrate input such
as potential used potential range, and plots at the bottom illustrates output,
such as current generated from a constant potential. (A) Chronoamperometry
(B) Cyclic Voltammetry (C) Linear Sweep Voltammetry.
x
2
2.1 Addition of FeCl2 enhances cathodic electron uptake by Shewanella oneidensis
MR-1. (A) Schematic of the bioelectrochemical reactor. The reactor consisted
of a stoppered glass bottle with a graphite working electrode (WE), platinum
counter electrode (CE), Ag/AgCl reference electrode (RE), and N2 ports. (B)
Chronoamperometry in anaerobic cathodic conditions with 30 mM fumarate
as the electron acceptor and the working electrode poised at +305 mV (vs
SHE) in the presence or absence of 1 mM FeCl2. 1 mM FeCl2 was added at
the time point indicated by the black arrow. Error bars indicate standard error
of triplicate measurements. (C) Representative cyclic voltammetry (1 mV/s
scan rate) of S. oneidensis with 30 mM fumarate as the electron acceptor in
the presence or absence of 1 mM FeCl2.
2.2 Cyclic voltammetry scan (-465 mV to 435 mV at 1 mV/s) of bare electrodes
with and without 1 mM FeCl2, in the presence of 30 mM fumarate. Cyclic
voltammetry reveals a quasi-reversible electron transfer behavior consistent
with the redox characteristics of FeCl2.
2.3 Representative fluorescence microscopy images of the graphite electrode with
attached WT S. oneidensis cells stained with FM 4-64FX cell membrane stain
with 1 mM FeCl2 (A) and without 1 mM FeCl2 (B). Images were taken after
15 h of staining. Fluorescence images shows similar cell density on the electrode
surface with or without exogenous FeCl2. This indicates that current
variability between conditions is due to the presence of soluble iron in the
system.
2.4 FeCl2 addition does not induce overexpression of heme-containing cytochrome
in S. oneidensis. Representative total cell protein SDS-PAGE and densitometry
analysis (A) Coomassie gel (B) TMBZ heme-stained SDS-PAGE gels.
2.5 The FeCl2-enhanced cathodic current into S. oneidensis is concentration
dependent. (A) Chronoamperometry (+305 mV vs SHE) of S. oneidensis with
increasing concentration of FeCl2: 0 mM, 0.2 mM, 0.6 mM and 1 mM. 30 mM
fumarate served as the electron acceptor. Error bars indicate standard error of
triplicate measurements. (B) Representative cyclic voltammetry scan (at 1
mV/s) of S. oneidensis with increasing concentration of FeCl2.
2.6 Fe-enhanced electron uptake by S. oneidensis MR-1 is diminished by the
addition of the iron chelator deferoxamine. (A) Representative
chronoamperometry curves of S. oneidensis on graphite electrodes poised at
+305 mV (vs SHE) with 30 mM fumarate as the electron acceptor. Arrow
indicates times of FeCl2 and DFO addition. (B) Representative cyclic
voltammogram profiles of cell-containing and control bare electrodes with 1
mM FeCl2 with and without deferoxamine.
xi
2.7 S. oneidensis MR-1 is not sensitive to DFO under the experimental conditions.
CFU counts via serial dilution before and after addition of 1.5 mM DFO.
2.8 Fe-enhanced electron uptake in S. oneidensis MR-1 is linked to fumarate
reduction. (A) Chronoamperometry measurement at +305 mV (vs SHE) of S.
oneidensis MR-1 WT and fumarate reductase mutant (∆fccA) on graphite
electrode surface in the presence of 1 mM FeCl2 and 30 mM fumarate as an
electron acceptor. Error bars indicate standard error of triplicate measurements. (B) Representative cyclic voltammogram of S. oneidensis MR-1 and
∆fccA on graphite electrode with 30 mM fumarate.
2.9 Cyclic voltammetry of S. oneidensis MR-1 with 1 mM FeCl2 without the
electron acceptor fumarate.
2.10 Fe-enhanced cathodic electron uptake is catalyzed by outer membrane
cytochromes in S. oneidensis MR-1. (A) Chronoamperometry experiments (-
305 mV vs SHE) of S. oneidensis, mtrC mutant (ΔmtrC), omcA mutant
(ΔomcA), mtrC and omcA double mutant (ΔmtrC/omcA), and outer
membrane cytochrome (OMC) mutant (ΔomcA/ΔmtrA/ΔmtrF/ΔdmsE/
ΔSO4360/ΔcctA/ΔrecA) with 30 mM fumarate serving as an electron acceptor. Addition of 1 mM FeCl2 is indicated by the arrowhead. Error bars indicate
standard error of triplicate measurements. (B) Representative cyclic
voltammograms of S. oneidensis WT and outer membrane cytochrome mutants
(+465 to 435 mV at 1 mV/s) under nitrogen atmosphere.
2.11 Representative fluorescence microscopy images of the graphite electrode with
attached S. oneidensis WT (A) and mutant cells stained with FM 4-64FX cell
membrane stain after electrochemical cultivation and cyclic voltammetry with
added FeCl2. Images were taken after 15 h of staining. Fluorescence images
shows similar cell density on the electrode surface with exogenous FeCl2. This
indicates that current variability between conditions is due to the availability
of heme containing cytochromes that shuttles electrons into the cell.
3
3.1 Graphical representation of the geographical location of DeMMO 4. Crosssectional view of the Deep Mine Microbial Observatory (DeMMO). Black lines
are tunnels and shafts in the mine. Pink circles represent locations of six
DeMMO sites, and larger circle represent location of DeMMO 4. Illustration
by Caitlin Casar (Casar et al., 2020).
3.2 Schematic diagram of parallel flow electrochemical reactor. Electrodes and
minerals were placed in glass chambers filled with water from the manifold.
Chambers were then submerged in water from the borehole to prevent direct
intrusion of ambient air entering reactors, while still allowing fluid flow.
Passive fluid flow from DeMMO 4 manifold was delivered to glass chambers,
and fluid escaped these chambers through a 7/8-inch diameter hole in the
xii
rubber stopper (not illustrated). The reference electrode (Ag/AgCl in 1 M KCl)
and counter electrodes (titanium wire) were submerged in glass reactor
effluent. A separate counter electrode was used for each channel, though only
one is illustrated for clarity. Duplicate ITO and graphite electrodes were poised
at ~-200 mV versus SHE, along with open circuit (OC) controls.
3.3 Pre-incubation linear sweep voltammograms (LSV) of filtered DeMMO4
fracture fluids using indium tin oxide (ITO). Scan rate at 0.1 mV/s within the
range of -600 mV to +500 mV (SHE).
3.4 Chronoamperometry profiles and cyclic voltammetry plots of Indium tin oxide
electrode 1 (ITO-1) for August-October incubation. (A) Chronoamperometry
profiles are shown for the 45-day incubation which were poised at ~−200 versus
SHE with data recorded at 30 s intervals. (B-F) Cyclic voltammetry plots ran
at a scan range of −600 mV to +300 mV versus SHE with a scan rate of 1
mV/s are also shown. Black circle symbols on chronoamperometry profile
correspond to the date in which the CV was performed: Sept. 8, Sept. 15, Sept.
22, Oct. 10, Oct. 16.
3.5 ITO-2 chronoamperometry profiles and cyclic voltammetry plots, (A)
Chronoamperometry profiles are shown for the 45-day incubation which were
poised at ~−200 versus SHE with data recorded at 30 s intervals. (B-F) Cyclic
voltammetry plots ran at a scan range of −600 mV to +300 mV versus SHE
with a scan rate of 1 mV/s are also shown. Black circle symbols on
chronoamperometry profile correspond to the date in which the CV was
performed: Sept. 8, Sept. 15, Sept. 22, Oct. 10, Oct. 16.
3.6 ITO-3 chronoamperometry profiles and cyclic voltammetry plots A)
Chronoamperometry profiles are shown for the 45-day incubation which were
poised at ~−200 versus SHE with data recorded at 30 s intervals. (B-F) Cyclic
voltammetry plots ran at a scan range of −600 mV to +300 mV versus SHE
with a scan rate of 1 mV/s are also shown. Black circle symbols on
chronoamperometry profile correspond to the date in which the CV was
performed: Sept. 8, Sept. 15, Sept. 22, Oct. 10, Oct. 16.
3.7 ITO-4 chronoamperometry profiles and cyclic voltammetry plots (A)
Chronoamperometry profiles are shown for the 45-day incubation which were
poised at ~−200 versus SHE with data recorded at 30 s intervals. (B-F) Cyclic
voltammetry plots ran at a scan range of −600 mV to +300 mV versus SHE
with a scan rate of 1 mV/s are also shown. Black circle symbols on
chronoamperometry profile correspond to the date in which the CV was
performed: Sept. 8, Sept. 15, Sept. 22, Oct. 10, Oct. 16.
3.8 Graphite 1 (GRA-1) chronoamperometry profiles and cyclic voltammetry plots
(A) Chronoamperometry profiles are shown for the 45-day incubation which
were poised at ~−200 versus SHE with data recorded at 30 s intervals. (B-F)
Cyclic voltammetry plots ran at a scan range of −600 mV to +300 mV versus
xiii
SHE with a scan rate of 1 mV/s are also shown. Black circle symbols on
chronoamperometry profile correspond to the date in which the CV was
performed: Sept. 8, Sept. 15, Sept. 22, Oct. 10, Oct. 16.
3.9 GRA-2 chronoamperometry profiles and cyclic voltammetry plots (A)
Chronoamperometry profiles are shown for the 45-day incubation which were
poised at ~−200 versus SHE with data recorded at 30 s intervals. (B-F) Cyclic
voltammetry plots ran at a scan range of −600 mV to +300 mV versus SHE
with a scan rate of 1 mV/s are also shown. Black circle symbols on
chronoamperometry profile correspond to the date in which the CV was
performed: Sept. 8, Sept. 15, Sept. 22, Oct. 10, Oct. 16.
3.10 GRA-3 chronoamperometry profiles and cyclic voltammetry plots (A)
Chronoamperometry profiles are shown for the 45-day incubation which were
poised at ~−200 versus SHE with data recorded at 30 s intervals. (B-F) Cyclic
voltammetry plots ran at a scan range of −600 mV to +300 mV versus SHE
with a scan rate of 1 mV/s are also shown. Black circle symbols on
chronoamperometry profile correspond to the date in which the CV was
performed: Sept. 8, Sept. 15, Sept. 22, Oct. 10, Oct. 16.
3.11 GRA-4 chronoamperometry profiles and cyclic voltammetry plots (A)
Chronoamperometry profiles are shown for the 45-day incubation which were
poised at ~−200 versus SHE with data recorded at 30 s intervals. (B-F) Cyclic
voltammetry plots ran at a scan range of −600 mV to +300 mV versus SHE
with a scan rate of 1 mV/s are also shown. Black circle symbols on
chronoamperometry profile correspond to the date in which the CV was
performed: Sept. 8, Sept. 15, Sept. 22, Oct. 10, Oct. 16.
3.12 Principal coordinate plot resulting from weighted-Unifrac PCoA analysis of the
resultant microbial communities based on OTUs. (A) Squares represent
samples from the May sampling trip, triangles represent samples from the
October sampling trip, and diamonds indicate filtered water samples taken
from the DeMMO 4 borehole. Legend colors indicate water sampling date,
graphite (GRA), indium tin oxide (ITO), sulfur (Sulf), pyrite and quartz source
materials. OC is used to indicated electrodes that were not poised, or open
circuit controls. Axis1 [35.2%] and Axis 2 [21.1%] indicate the percent variation
explained. (B) Principal coordinate plot of the weighted-Unifrac PCoA analysis
of the October in situ microbial community samples. Squares represent
graphite samples (GRA), triangles represent ITO samples, diamonds represent
mineral samples (sulfur [Sulf], pyrite, and quartz [Qrtz]) and the plus sign
indicates filtered water sampled from the DeMMO 4 borehole. Axis 1 [30.3%]
and Axis 2 [16.8%] signify the percent variation explained.
3.13 Relative 16S rRNA abundances for all SURF community samples collected at
the order level. Orders for the top 100 OTUs plotted for clarity. Remaining
OTUs were less than 0.13% abundance of total sequences.
xiv
4
4.1 A. actinomycetecomitans produces soluble electron shuttles.
Chronoamperometry measurement on A. actinomycetecomitans with the
working electrode poised at +435 mV (vs SHE) with 30 mM DL-lactate. Arrow
heads indicate inoculation time, fresh media exchange and reintroduction of
spent media. Reactors were kept in nitrogen atmosphere at 37 °C.
4.2 A. actinomycetecomitans robust biofilm on the electrode. Scanning Electron
Microscopy (SEM) of A. actinomycetecomitans biofilm on graphite felt
electrode fibers.
4.3 Scan rate dependence of the cyclic voltametric response of spent media A.
actinomycetecomitans. (A) Cyclic voltammetry measurement of reactor spent
media. The potential was swept -700 mV to 200 mV vs SHE at 1 mV/s, 10
mV/s, 50 mV/s and 100 mV/s. inset shows voltammetry measurements of
sterile media. (B) Peak current vs scan rate plot.
4.4 Addition of FMN does not improve anodic current relative to the negative
control. Chronoamperometry measurement on A. actinomycetecomitans with
the working electrode poised at +435 mV (vs SHE). Arrow heads indicate
addition of 200 nM and 400 nM FMN. Reactors were kept in nitrogen
atmosphere at 37 °C.
4.5 A. actinomycetecomitans is not sensitive to cytochrome inhibitors KCN and
NaN3. Chronoamperometry measurement on A. actinomycetecomitans with
the working electrode poised at +435 mV (vs SHE). Arrow heads indicate
addition of 5 mM KCN and 10 mM NaN3. Reactors were kept in nitrogen
atmosphere at 37 °C.
4.6 Current levels between Live and Heat-killed A. actinomycetecomitans are
comparable. Average anodic current levels between Live and Heat-killed cells.
Error bars represent standard error of triplicate experiments. Working
electrode poised at +435 mV (vs SHE) with 30 mM DL-lactate. Reactors were
kept in nitrogen atmosphere at 37 °C.
4.7 A. actinomycetecomitans generates anodic current lower than S. oneidensis
∆OMC mutant. Chronoamperometry measurements on WT S. oneidensis MR1, outer membrane cytochrome (OMC) mutant (ΔomcA/ΔmtrA/ΔmtrF/
ΔdmsE/ΔSO4360/ΔcctA/ΔrecA), and A. actinomycetecomitans (Aa) with 30
mM DL-lactate as the sole electron source. WE poised at +435 mV (vs SHE)
under nitrogen atmosphere.
xv
List of tables
Table 2.1. Strains used to study the role of S. oneidensis outer membrane
cytochromes in FeCl2-enhanced extracellular electron uptake.
Table 3.1 Geochemical measurements from DeMMO 4 at SURF, before and after
electrochemical cultivation.
Table 3.2 Significantly enriched OTUs on ITO electrodes compared with ITO
open circuit controls as assessed by DEseq2.
xvi
Abstract
Extracellular electron transfer (EET) is a respiratory strategy that allows bacteria to
access external electron acceptors such as redox-active elements (e. g., Fe, Mn) in solid
minerals. Extracellular electron uptake (EEU), on the other hand, refers to the capacity
of microorganisms to acquire electrons from solid-phase conductive materials like metal
oxides, a less understood mechanism. Organisms such as Shewanella oneidensis and
Geobacter sulfurreducens are two of the well-studied organisms capable of EET. In S.
oneidensis, the oxidation of a carbon source occurs in the cytoplasmic space and gained
electrons are transferred from heme containing inner membrane proteins, through the
periplasm and eventually to the outer membrane reducing external electron acceptors. To
date there are three known ways S. oneidensis respire external electron acceptors, by direct
contact of outer membrane cytochromes, through soluble electron shuttles, and lastly via
membrane extensions containing outer membrane cytochromes. Much of our mechanistic
understanding of EET is derived from studies of transmembrane cytochrome complexes
and extracellular redox shuttles that mediate outward EET to anodes and external
electron acceptors. In contrast, there are knowledge gaps concerning the reverse process
of inward EET (EEU) from external electron donors to cells. To address this knowledge
gap, the first project discussed in this dissertation touches on the role of soluble iron in
xvii
enhancing EET from cathodes to the model EET bacterium S. oneidensis MR-1. Here we
use amperometric and voltammetric tools coupled with gene deletions to demonstrate that
S. oneidensis can use FeCl2 as an electron shuttle to enhance cathodic electron uptake.
Our data reveals that FeCl2 does enhance electron uptake by S. oneidensis with the
involvement of the Mtr pathway all the way to the periplasm reducing electron acceptor
fumarate to succinate.
The second project we discuss in this dissertation looks into how EET may
contribute in maintaining microbial communities in the deep-subsurface terrestrial
environments. At the Deep Mine Microbial Observatory (DeMMO) located in Sanford
Underground Research Facility (SURF) in Lead, South Dakota, we conducted long-term
in situ electrochemical experiments aimed at evaluating the capability of deep
groundwater microbial communities to employ extracellular electron transfer to support
microbial respiration. We couple electrochemical measurements with 16S rRNA analysis
to look at the influence of poised electrodes on the microbial population on a conductive
surface. Our results indicate that there are anodic and cathodic activities at the site.
Moreover, the ability to attach to surfaces is the main driver of microbial composition on
minerals and electrodes.
In the last chapter of this dissertation, we focus on the human microbiome where
the number of bacteria that are known to be capable of EET is relatively small compared
to those that are found in the environment. Specifically, we performed electrochemical
xviii
investigations on a microbial isolate from the oral microbiome, Aggregatibacter
actinomycetecomitans, previously proposed to be capable of EET. Aggregatibacter
actinomycetecomitans D7S-1 represents a significant threat to human health, being a
Gram-negative opportunistic pathogen and a member of the oral microbiota strongly
associated with severe types of periodontal disease. Amperometric and inhibition assays
were conducted to investigate the extent of microbial electrochemical activity of this
strain. While the inoculation of electrochemical reactors with A. actinomycetecomitans
results in weak current production, our study highlights the importance of controls
designed to test whether the observed current in such ‘weak electrogens’ stems from in
vivo activity.
In sum, we discovered an inward Fe-mediated extracellular electron transfer (EET)
mechanism in a well-studied EET model organism marked a significant breakthrough.
Additionally, we expanded electrochemical studies to include “non-canonical” EET
microbes, both in their natural environment and within the human microbiome. This
expansion broadened our understanding of microbial electrochemical interactions and
their potential roles in various ecosystems and human health.
1
Chapter 1: Introduction
1.1 Microbiological perspective of biodiversity
Before the great oxygenation event, the Earth’s atmosphere, largely devoid of molecular
oxygen, underwent a pivotal transformation due to the evolution of oxygenic
photosynthesis by ancient cyanobacteria. (Staley & Reysenbach, 2002) The increase in
atmospheric oxygen had profound effects on early life forms, paving the way for the
evolution of more complex organisms that relied on aerobic respiration. However, many
of the early bacteria were anaerobic or facultative, that is, they are able to grow in
environments in the presence or absence of oxygen. Anaerobic bacteria thrived in early
Earth environments due to their ability to use various metabolic pathways that do not
require oxygen, such as anaerobic respiration, fermentation, and chemosynthesis to obtain
energy from organic and inorganic compounds. (Kirchman, 2018; Staley & Reysenbach,
2002) Nevertheless, O2 was toxic to many anaerobic organisms, over time, this resulted in
evolution of aerobic metabolism. Organisms that developed mechanisms to tolerate or
utilize oxygen were more likely to thrive in the newly oxygen-rich environment, ultimately
leading to the diversification of life forms and development of complex ecosystems.
2
Anaerobic bacteria played a critical role in shaping early Earth environment and
the evolution of life. These organisms shape biogeochemical cycles and mediate the cycling
of essential elements such as carbon, nitrogen, sulfur and iron. (Perry, 2002; Staley &
Reysenbach, 2002) For example, sulfate-reducing bacteria play an essential role in the
sulfur cycle by reducing sulfate to hydrogen sulfide and other sulfur species. (Agostino &
Rosenbaum, 2018; Guan et al., 2015) One of the mechanisms employed by methanogenic
bacteria involves converting carbon dioxide and hydrogen into methane to generate energy
for the cells. (Enzmann et al., 2018; Hara et al., 2013; Rowe et al., 2019) Denitrifying
bacteria can use nitrate as an electron acceptor. (Gregoire et al., 2014; Pous et al., 2014)
Iron reducing bacteria use iron oxides as terminal electron acceptors similar to eukaryotic
organisms use oxygen. (Nealson & Saffarini, 1994) The dynamic nature of ecosystems was
influenced by the metabolic adaptability and ecological roles of microorganisms, which
paved the way for subsequent evolutionary advancements. In this dissertation, we explore
the microbial interactions with redox-active minerals or electrodes, with a specific
emphasis on organisms capable of utilizing electrodes as external sources of electron
acceptors or donors for energy production.
1.2 Oxidative phosphorylation
All extant life requires energy for biosynthetic processes, essential for synthesizing complex
molecules including nucleic acids, proteins, carbohydrates, and lipids. (Amend et al., 2013;
Haddock & Jones, 1977; Kirchman, 2018; Staley & Reysenbach, 2002) Cellular electron
3
transport in particular is crucial for energy production in most living organisms. To delve
into the intricacies of electron transport, it is paramount to first discuss aerobic
respiration, that is, oxidative phosphorylation. (Hatefi, 1985)
Oxidative phosphorylation is a cellular process that harnesses the reduction of
oxygen to generate high-energy phosphate bonds in the form of adenosine triphosphate
(ATP). (Figure 1.1) During oxidative phosphorylation, electrons derived from oxidation
of an electron source such as organic carbons are transferred along a series of electron
Figure 1.1. Schematic representation of oxidative phosphorylation in the mitochondria.
Electrons transferred through the electron transport chain in the inner mitochondrial
membrane, generating a proton gradient that drives ATP synthesis by ATP synthase.
This process is crucial for cellular energy production in aerobic respiration. Illustration
from (Shrestha, 2003). This electron transport chain is composed of complex I
(NADH:ubiquinone oxidoreductase) complex II (succinate:ubiquinone oxidoreductase),
coenzyme Q10 (CoQ), complex III (ubiquinol:cytochrome c oxidoreductase), cytochrome
C, complex IV (cytochrome c oxidase), and complex V (H+-translocating ATP synthase).
4
transfer proteins embedded in the inner mitochondrial membrane to reduce oxygen and
form water. The flow of electrons releases energy that allows for the transport of protons
from the inner membrane to the periplasm, establishing an electrochemical gradient
characterized by higher proton concentration in the periplasmic space. The
electrochemical gradient established then serves as the driving force for ATP synthesis.
The synthesis of ATP from adenosine diphosphate (ADP) and inorganic phosphate (Pi)
is catalyzed by the flow of protons back into the mitochondrial cytoplasmic space through
the ATP synthase complex. (Hatefi, 1985) In early Earth, however, free oxygen was not
available. Microorganisms therefore have adapted to derive energy from oxidizing
compounds by evolving the capacity to utilize alternative electron acceptors in addition
to oxygen. Hence, organisms capable of using electron acceptors other than oxygen were
the dominant population. (LaRowe & Amend, 2015; Myers & Nealson, 1988; M. R. Osburn
et al., 2014)
Substances differ in their tendency to undergo oxidation or reduction reactions. By
convention we describe redox reactions with respect to the reduction potential (Eo’,
standard conditions). This term quantitatively describes the ability of an element to gain
electrons. A variety of compounds can participate in redox reactions. (Amend & Shock,
2001; Madigan & Martinko, 2006; Osburn et al., 2014) Under varying conditions, these
compounds can function as either electron donors or electron acceptors, depending on the
substance with which they react. A practical approach to understand electron transfer
5
reactions in biological systems and their significance in bioenergetics is to envision a
vertical tower. The Redox Tower graphically tabulates redox half reactions based on their
Eo’ values, to predict the direction of electron flow between potential electron donors and
acceptors. (Figure 1.2)
Figure 1.2. Schematic of redox tower showing electron donors and acceptors used by
microorganisms. Electron donors and acceptors are arranged with respect to their standard
reduction potential (standard conditions). Electron donors with a more reducing potential
are at the top of the tower while the electron acceptors with a more oxidizing potential
are localized at the bottom. Arrows indicate electron donor/acceptor pairs that certain
microbes prefer with respect to their metabolic machinery. Figure courtesy of Moh ElNaggar.
6
The Redox Tower ranks compounds from the most negative Eo’, compounds that
can readily lose electrons, to the most positive Eo’, compounds that will most likely accept
electrons. For example, O2 is a very strong oxidizing agent that electrons will naturally
gravitate towards, placing O2 at the bottom of the redox tower with the most positive Eo’
out of all electron acceptors. The redox tower then can help infer the electron acceptor
preference of the microbes based on the electron donors available. Microorganisms can
utilize a variety of electron acceptors during anaerobic respiration with respect to their
metabolic capabilities and the environmental conditions. This, in turn, allows them to
adapt to changing environmental conditions.
1.3 Extracellular electron transfer
Extracellular electron transfer is the process by which bacteria use an electron acceptor
or an electron source located outside of the cell. (Barco et al., 2015; Chong et al., 2018;
Emerson et al., 2010; Myers & Nealson, 1988; Summers et al., 2013; Zacharoff & ElNaggar, 2017) Two of the of organisms that are known to interact with minerals outside
of the cell for respiration are Shewanella oneidensis and Geobacter sulfurreducens. (Marsili
et al., 2010; Xu et al., 2016) S. oneidensis and G. sulfurreducens can reduce manganese
7
Figure 1.3. Crystal structure of the outer membrane metal reducing (mtr) pathway in S.
oneidensis MR-1. Crystal structure and heme arrangement of extracellular facing outer
membrane cytochrome MtrC (blue). Crystal structure and heme arrangement of outer
membrane-associated periplasmic cytochrome MtrA (pink). Crystal structure of beta
barrel MtrB (green). Figure obtained from (Edwards et al., 2020).
and iron oxides and electrodes as electron acceptors. Since the discovery of these two
organisms, it has been revealed that this ability to interact with the “outside world” is
mediated by proteins spanning the outer membrane of the cell. These proteins contain
hemes with iron centers which act as a wire that transports current (movement of
electrons) from inside the cell to outside of the cell. (Figure 1.3) (Clarke et al., 2011;
Edwards et al., 2018, 2020) For S. oneidensis, electrons start at the cytoplasmic region
where the electron is oxidized to NADH/MQ oxidoreductase passing electrons to the
quinol pool where the reduced quinones are oxidized by the inner-membrane associated
8
tetraheme menaquinone dehydrogenase CymA. Periplasmic multiheme cytochromes (STC
and FccA) get reduced by CymA, transferring electrons to the outer membrane bound
MtrABC porin-cytochrome complex. (Figure 1.4) MtrA is a decaheme c-Cyt that is
embedded in MtrB, a trans outer membrane porin-like protein. MtrAB facilitates electron
transfer to the extracellular decaheme c-Cyt MtrC and OmcA from which extracellular
electron acceptors are reduced. (Breuer et al., 2014; Edwards et al., 2020; Richardson et
al., 2012; White et al., 2016) In addition to previously mentioned protein-protein
interactions, the outer membrane proteins OmcA and MtrC interact with each other,
facilitating electron transfer on the cell surface. (Chong et al., 2022)
Currently, S. oneidensis is known to perform EET in three ways: (1) direct electron
transfer, where the cell is directly attached to the substrate allowing for the outer
membrane cytochromes to donate electrons to the electron acceptor; (Breuer et al., 2014;
Coursolle & Gralnick, 2012) (2) redox shuttle mediated, here the presence of electron
carriers such as flavins allows the cells to get “rid” of the electrons; (Okamoto, Kalathil,
et al., 2014; Xu et al., 2016) (3) and lastly, through nanowires embedded with outer
membrane cytochromes, which facilitates electron transfer over relatively long distances.
(Figure 1.4) In this case, the cell is not directly attached to the electron acceptor but an
“appendage”, the nanowire, is in contact and connects the cell to the electron acceptor.
(El-Naggar et al., 2010)
9
Figure 1.4. Schematic representation of extracellular electron transfer in dissimilatory
metal reducing bacteria Shewanella oneidensis MR-1. Electron transfer to the extracellular
space facilitated by inner membrane and periplasmic redox-active proteins and is handed
off to outer membrane multiheme cytochromes. Extracellular electron transfer to solid
surfaces is enabled by outer membrane cytochromes directly interacting with solid surface,
cytochrome bound cofactors, and nano-wire also known as membrane extensions consisting
outer membrane cytochromes or indirectly via soluble electron shuttles. Figure obtained
from (Chong et al., 2018)
10
1.4 Extracellular electron uptake
Extracellular electron transfer (EET) extends beyond outward (cell-to-surface)
interactions. Several microorganisms, including iron (II)-oxidizing bacteria can obtain
electrons from outside of the cell and derive energy through inward EET. (Bose et al.,
2014; Ross et al., 2011; Rowe et al., 2018) This capability is essential for microbial
electrosynthesis technologies, where EET from cathodes facilitates the reduction of CO2,
enabling the production of biofuels or other chemicals. However, the underlying
mechanisms of this cathodic extracellular electron uptake remain less understood than its
anodic counterpart, but also include mediated EET via small molecules (e. g., H2) and
direct EET via porin-cytochrome complexes analogous to those described in Shewanella.
There are a three known mechanisms for electron uptake, (1) through external
hydrogenases, (2) through abiotic production of hydrogen, and (3) via direct interaction
of cytochrome to the metal (liquid or solid phase). (Aryal et al., 2017; Deutzmann et al.,
2015; Jain et al., 2022; Ross et al., 2011; Rowe et al., 2018; Summers et al., 2013) (Figure
1.5) Hydrogenases are enzymes that can catalyze the oxidation and production of
molecular hydrogen. Bacterial hydrogenases play a crucial role in acting as a mediator for
electron uptake from H2. Organisms such as Methanococcus maripaludis for example
specialize in hydrogen oxidation. (Deutzmann et al., 2015) On a cathode, hydrogen can
also interact directly with the electrode following the reaction H2 → 2H+ + 2e-
. Certain
bacterial species can capitalize on this and take up the electrons generated from this
11
Figure 1.5. Proposed mechanisms of extracellular electron uptake from a cathode. (A)
mediated electron transfer (B) mediated electron transfer via hydrogenase generated H2
(C) Direct electron transfer through outer membrane cytochromes.
reaction. (Aryal et al., 2017) Other organisms capable of electron uptake use c-type
cytochromes on the outer membrane of the cell. Rhodopseudomonas palustris TIE-1 can
take up electron via the photosynthetic electron uptake pathway PioABC. (Bose et al.,
2014) TIE-1 can utilize a variety of electron donors, including iron and cathodes. (Gupta
et al., 2020; Guzman et al., 2019) This three-component electron transport system moves
electrons from PioA, which sits in the porin structure PioB, and transfers electrons to
PioC from which electrons gained from the cathode or iron enters the quinone pool.
Siderooxydans lithotrophicus ES-1, Thioclava electrotropha ElOx9, and Mariprofundus
ferrooxidans are some of the other organisms known to perform EEU. (Jain et al., 2022;
Karbelkar et al., 2019; Summers et al., 2013).
While S. oneidensis is known for its capability to respire anodes and metal oxides,
it has also been investigated for its ability to take up electrons from a cathode. (Abuyen
12
& El-Naggar, 2023; Ross et al., 2011; Rowe et al., 2018) Investigations on S. oneidensis
are enticing given its genetic tractability and that the organism is easily culturable. To
that end several researchers have directed their investigation on S. oneidensis and its
ability to take up electrons. It has been shown that cathodic cultivation of S. oneidensis
leads to an increase its reducing equivalents during cathodic conditions. (Ross et al., 2011;
Rowe et al., 2018) In the second chapter we describe the role for soluble iron (exogenous
FeCl2) in augmenting extracellular electron transfer from cathodes to the model EET
bacterium S. oneidensis MR-1.
1.5 Microbial electrochemistry
Microbial electrochemistry is an interdisciplinary field that converges microbiology,
electrochemistry, and environmental science, which explores the interactions between
microorganisms and electrodes. In recent years, there has been an increasing focus on
understanding and leveraging microbial electrochemical processes for various applications,
ranging from environmental remediation to bioenergy production. At the heart of a single
chamber electrochemical reactor is a working electrode (WE), a counter electrode (CE),
and a reference electrode (RE) (Figure 1.6). The WE is where the bacteria will interact
and where the potential or voltage is set to mimic the redox potential of a particular metal
that can be reduced by the bacteria. The CE has the purpose of completing the circuit to
allow charge flow. Consequently, the material at which the CE is manufactured must be
inert such as platinum or carbon. Moreover, the size the CE needs to be larger
13
Figure 1.6. Schematic representation of bioelectrochemical reactors. (A) The reactor
consists of a glass cylinder attached to a working electrode (WE) (gold or ITO-coated
glass coverslip) at the base. The top is sealed with a cap that holds the counter electrode
(CE) (Pt wire), the reference electrode (RE), and the nitrogen ports. (B) The reactor with
a 3D WE (graphite).
than the WE to prevent current limitation which becomes more of an issue when the
organisms interacting with the electrode generates a lot of current. Lastly, the reference
electrode (RE) provides a reference potential and allows for measuring the change in
potential difference between the WE and CE. Silver/silver chloride (Ag/AgCl) is a
common reference electrode choice for biological electrodes. The electrode functions as a
redox electrode and the reaction is between the silver metal (Ag) and salt–silver chloride
(AgCl). The corresponding reaction can be observed as Ag
+ + 1e
- → Ag
O(s).
Common electrochemical measurements include various techniques that are used
to study the behavior of electrochemical systems and processes. Some of the
14
electrochemical measurements used in this dissertation include cyclic voltammetry,
chronoamperometry, and linear sweep voltammetry. Cyclic voltammetry is a versatile
technique used to investigate the redox behavior of an electroactive species in solution.
This technique involves applying a potential sweep to the working electrode while
measuring the current. (Figure 1.7) The scan rate has a significant influence on the shape
and characteristics of the resulting voltammogram. As the scan rate increases, the peak
current magnitude generally increases. This is because the faster scan rate shortens the
time for the electrochemical reactions to occur at the electrode surface at a particular
potential. By varying the scan rate, researchers can gain insights into the kinetic and
diffusion-controlled processes occurring at the electrode interface and extract valuable
information. (Elgrishi et al., 2018)
Chronoamperometry involves measuring the current at a constant potential,
typically in response to the electrochemical reaction of an analyte at the working electrode
surface. (Figure 1.7) This technique is usually used for continuous monitoring of an analyte
or electroactive bacteria. Several conditions can affect chronoamperometry measurements.
The surface area of the electrode directly influences the current response in
chronoamperometry, where a larger surface area allows for more electroactive species to
interact with the electrode and therefore resulting in higher currents. Temperature can
also affect the kinetics of electrochemical reactions and the diffusion of electroactive
species to the electrode surface. Changes in temperature may alter reaction rates, diffusion
15
Figure 1.7. Schematic of the electrochemical techniques used in this study. Top figures
illustrate input such as potential (E) used or potential range, and plots at the bottom
illustrates output, such as current (I) generated from a constant potential over time (t).
(A) Chronoamperometry measurement where a constant potential is set on the WE and
current is monitored over time. (B) Cyclic Voltammetry is when we scan a particular
range of potentials at a particular scan rate and measure the current generated throughout
the scan, forward and reverse. (C) Linear Sweep Voltammetry is similar to Cyclic
voltammetry in that current is measured over a particular potential range in one direction.
coefficients, and mass transport properties which ultimately leads to variations in the
current response. Changes in pH may also alter the redox species especially if the species
being investigated is biological. That said, it is important to ensure that temperature and
pH remains constant throughout the whole experiment to have reproducible results. The
presence of interfering species in solution can also impact the measurement as this will be
affecting the selectivity of the electrochemical measurement. Interfering species may
compete with the target analyte for adsorption leading to changes in the observed current.
16
Linear sweep voltammetry is where a constant rate of potential sweep is applied to
the working electrode using a potentiostat. (Figure 1.7) The voltammogram obtained from
this technique provides valuable information about the redox behavior of the analyte.
Peaks in the voltammogram correspond to electrochemical reactions occurring. Position,
magnitude, and shape of these peaks can provide information on the electrochemical
kinetics, redox potentials and concentration of electroactive species.
The use of electrochemical reactors has become an important tool in understanding
cellular electron transfer by microorganisms in the environment. (Jangir et al., 2016; Lam
et al., 2019; Rowe et al., 2014) Electrochemical techniques have their advantages due to
their simplicity (miniature and portable) and low economic burden compared to other
analytical techniques, while providing sensitivity and selective analysis. This allows
investigators to control activity and redox potential in the environment in which the
bacteria lives. (Jangir et al., 2016; Lam et al., 2019; Rowe et al., 2014) More importantly,
researchers are using electrochemical setups as an isolation tool that allows them to select
for organisms that can reduce or oxidize metal oxides. (Jangir et al., 2016; Karbelkar et
al., 2019; Rowe et al., 2014)
1.6 Microbes in anoxic environments
While there is a diversity of bacterial species there is likewise a diverse set of environments
within which these organisms can be found. Strains have been isolated from deep-sea
17
environments, in the sediment and areas where hydrothermal vents spew nutrients and
metals in the surrounding area for the native bacteria to metabolize. (Emerson & Moyer,
2002; Pérez-Rodríguez et al., 2021; Smith et al., 2018; Tully et al., 2017) In terrestrial
environments, metal reducing bacteria are also found in lakes, deep subsurface, and desert
to name a few. (Jangir et al., 2016; Karbelkar et al., 2019; Pérez-Rodríguez et al., 2021)
Subsurface environment below the ocean sediments and soil is largely devoid of oxygen.
(LaRowe & Amend, 2015; Staley & Reysenbach, 2002) In sediments, aerobic heterotrophy
consumes oxygen fast that it prevents oxygen from penetrating far, thus creating anoxic
mud only millimeters away from the surface. (Emerson et al., 2010; Kirchman, 2018)
Oxygen deficient environments have varying sizes and shape. Large or small however,
anoxic environments are dominated by archaea and bacteria. (Amend & Shock, 2001;
Author et al., 1996)
In addition to handling environments devoid of oxygen, living organisms in the
terrestrial subsurface must be able to maintain a dormant state for long periods of time
or metabolize and reproduce using resources found in situ. Energy resources available are
those transported to the subsurface by the hydraulic cycle and the rock cycle. (Kirchman,
2018; Staley & Reysenbach, 2002) Sedimentation and burial transports organic matter
from the surface biosphere to the subsurface. Percolation of groundwater carries gas into
the subsurface. Volcanism moves reduced minerals and gas from the Earths mantle to the
biosphere. Additionally, several sources of H2 are present in the subsurface which serves
18
as the electron donor for microbial primary production. Due to the vast and heterogeneous
nature of the terrestrial subsurface, it is challenging to get a precise estimation of the total
biomass. It was only recently that that we have come to recognize that these habitats
might constitute the most extensive biosphere on Earth, both in terms of the quantity of
the cells and the overall biomass. (Bar-On et al., 2018; Magnabosco et al., 2018)
Nevertheless, there is great potential for studying these kinds of environments.
Consequentially, we wanted to investigate the microbe-mineral interactions in the
subsurface environments to investigate the role of extracellular electron transfer in this
environment. In the 3rd chapter we show how electrochemical techniques can be useful in
investigations in subsurface environments. Here we discuss long-term investigation using
electrodes and minerals at Sanford Underground Research Facility (SURF) in South
Dakota to study microbe-mineral interactions.
1.7 The Human Microbiome
The human microbiome plays a critical role in maintaining human health via contributions
to digestion, nutrient absorption, and immune system development. Consequently,
changes in the composition and function of the microbiome have been linked to diseases
such as gastrointestinal disorders, metabolic disorders, and even autoimmune conditions.
(Carbonero et al., 2012; Wanger et al., 2013) Several species have this critical role in the
human gut. Bacteriodes fragilis and Prevotella spp., are important members of the gut
microbiota as they are involved in the fermentation of complex carbohydrates and
19
production of short-chain fatty acids. (Flint et al., 2012; Zafar & Saier, 2021) Akkermansia
muciniphila, is a mucin-degrading bacterium that resides in the mucus layer of the gut
epithelium and plays a critical role in maintaining gut barrier integrity. (Glover et al.,
2022) Escherichia coli, while some strains are pathogenic, others are commensal bacteria
that contribute to the production of vitamins in addition to colonization resistance against
pathogens. (Bentley & Meganathant, 1982; Lawrence & Roth, 1996)
In the human microbiome, organisms capable of extracellular electron transfer have
been identified, although their presence and function have yet to be extensively
investigated as in other environments such as aquatic or soil ecosystems. Pathogens that
have caused serious illnesses have been found to be electrochemically active. Listeria
monocytogenes, a Gram-positive bacterium known to cause listeriosis, has been shown to
be capable of extracellular electron transfer by producing flavins. (Light et al., 2018)
Enterococcus faecalis, another Gram-positive bacterium found in the human
gastrointestinal tract likewise has the ability to perform flavin-mediated EET. (Keogh et
al., 2018) Surprisingly, Listeria monocytogenes and Enterococcus faecalis have similar
current levels as S. oneidensis. (Keogh et al., 2018; Light et al., 2018)
The electrochemical activity of Listeria monocytogenes was discovered several
years ago, but only recently has research emerged regarding the organism’s utilization of
flavin-mediated extracellular electron transfer. (Deneer & Boychuk, 1993; Light et al.,
2018) An eight-gene locus was discovered to govern the organism’s capacity to perform
20
EET, involving a process where NADH dehydrogenase channels electrons to a quinone
pool localized in the membrane, in conjunction with an extracellular flavo-lipoprotein
where flavins are facilitating electron transfer to extracellular electron acceptors. Given
the significance of anaerobic growth mechanisms in microbial proliferation within the
intestinal lumen, it has been hypothesized that L. monocytogenes (and other food borne
pathogens) might employ EET to thrive in the gut, particularly since the activation of
EET can support growth in the presence of non-fermentable carbon sources. (Light et al.,
2018)
Organisms from the oral microbiome have been proposed to be electrochemically
active as well. In the oral microbiome, bacteria contribute to various physiological
processes and influence oral health and disease. While bacteria can be associated with oral
disease, there are also members of the oral microbiome that positively contribute to oral
health. Beneficial bacteria like Streptococcus salivarius and Lactobacillus species aid in
oral homeostasis by competing with pathogenic bacteria and producing antimicrobial
compounds. (Khalfallah et al., 2021) Other key roles of bacteria in the oral microbiome
include: dental plaque formation, fermentation and acid production, influence on
periodontal health and disease, some contribute to immune regulation and homeostasis in
the oral environment. It is still true however that bacteria are intimately involved in the
development of periodontal disease. This imbalance in the oral microbiome can lead to
inflammation of the gums and deconstruction of periodontal tissues. (Loesche &
21
Grossman, 2001; Tian et al., 2010) One of the organisms that are associated with
periodontal disease is Aggregatibacter actinomycetecomitans. A. actinomycetecomitans is
of particular interest to researchers due to its strong association with aggressive
periodontitis, a severe form of periodontal disease that is characterized by rapid loss of
periodontal attachment and bone destruction. More importantly, A.
actinomycetecomitans is considered the key pathogen driving disease progression.
Scientific work has suggested that A. actinomycetecomitans is capable of extracellular
electron transfer. (Naradasu et al., 2020) In collaboration with Casey Chen’s lab from
USC School of dentistry, we further investigated the EET capability of A.
actinomycetecomitans, which will be discussed in Chapter 4.
Studying the human microbiome has far-reaching implications for health, medicine
and the environment. By investigating the interactions between the human host and
microbial communities, we can gain valuable insights into health and disease paving the
way for development of innovative approaches for promoting wellness and preventing
illness. EET-capable microorganisms can be utilized in electrochemical sensors and
biosensors for detection and monitoring. (Atkinson et al., 2023) By harnessing the
sensitivity and specificity of the bacterial EET process, these sensors can then offer rapid,
cost-effective and portable solutions for medical diagnostics.
22
Chapter 2: Soluble Iron Enhances
Extracellular Electron Uptake by
Shewanella oneidensis MR-1
This chapter has been adapted from (Abuyen & El-Naggar, 2023):
Abuyen, K., & El-Naggar, M. Y. (2023). Soluble Iron Enhances Extracellular Electron
Uptake by Shewanella oneidensis MR-1. ChemElectroChem, 10(4).
DOI: 10.1002/CELC.202200965. https://doi.org/10.1002/celc.202200965
23
2.1 Abstract
Extracellular electron transfer (EET) is a process that microorganisms use to reduce or
oxidize external electron acceptors or donors in solid phase. Much of our mechanistic
understanding of this process is derived from studies of transmembrane cytochrome
complexes and extracellular redox shuttles that mediate outward EET to anodes and
external electron acceptors. In contrast, there are knowledge gaps concerning the reverse
process of inward EET from external electron donors to cells. Here, we describe a role for
soluble iron (exogenous FeCl2) in enhancing EET from cathodes to the model EET
bacterium Shewanella oneidensis MR-1, with fumarate serving as the intracellular electron
acceptor. This iron concentration-dependent electron uptake was eradicated upon addition
of an iron chelator and occurred only in the presence of fumarate reductase, confirming
an electron pathway from cathodes to this periplasmic enzyme. Moreover, S. oneidensis
mutants lacking specific outer membrane and periplasmic cytochromes exhibited
significantly decreased current levels relative to wild-type. These results indicate that
soluble iron can function as an electron carrier to the EET machinery of S. oneidensis.
2.2 Introduction
Some microorganisms, such as the metal-reducing bacterium Shewanella oneidensis MR1, are capable of performing extracellular electron transfer (EET), a respiratory strategy
that allows them to access external electron acceptors such as redox-active elements (e.g.
24
Fe, Mn) in solid minerals. (Myers & Nealson, 1988; Shi et al., 2016) Importantly, this
process can also be exploited in bioelectrochemical technologies, such as microbial fuel
cells, and living electronics that ‘wire’ microbes to anodes. (Abbas et al., 2017; Nealson,
2017)
In the case of S. oneidensis, the challenge of outward EET is overcome by a network
of cytochromes (collectively referred to as the Mtr pathway) that bridge the otherwise
electrically insulating cell envelope. This network includes the inner membrane tetraheme
cytochrome CymA, periplasmic cytochromes including the small tetraheme cytochrome
(CctA) and flavocytochrome fumarate reductase FccA, the transmembrane porincytochrome complex MtrAB, and the cell surface decaheme cytochromes MtrC and OmcA
that function as electron conduits to the outside world. (Coursolle & Gralnick, 2010, 2012;
Ross et al., 2011) When these cell surface conduits transfer electrons to external surfaces
(e.g. minerals or electrodes) through direct contact, this process is referred to as direct
EET. EET can also proceed to external surfaces indirectly via soluble diffusing redoxactive shuttles, such as flavins. (Coursolle et al., 2010; Okamoto, Nakamura, et al., 2014)
Flavins can also play a role as cytochrome-bound cofactors that enhance EET rates. (Xu
et al., 2016) Interestingly, there is evidence that soluble iron may also serve as an
extracellular mediator linking the cellular EET machinery to anodes by redox cycling.
(Oram & Jeuken, 2016)
25
EET is not only an outward (cell-to-surface) phenomenon. A number of
microorganisms, such as Fe(II) oxidizing bacteria, can acquire electrons from the exterior
of the cell and derive energy from this inward EET. (Carbajosa et al., 2010; Karbelkar et
al., 2019; Summers et al., 2013) Other microbes are proposed to utilize direct electron
uptake, such as corrosive sulfate reducing bacteria. (Hamilton, 2003) For example, the
sulfate reducing bacteria such as Desulfovibrio ferrophilus IS5 and Desulfopila corrodens
IS4 have been shown to directly interact with electrodes and demonstrated electron uptake
from cathodes. (Beese-Vasbender et al., 2015; Deng et al., 2015; McCully & Spormann,
2020) This ability is crucial for microbial electrosynthesis technologies, where EET from
cathodes drives reduction of CO2 and production of biofuels or other chemicals. (Hara et
al., 2013; Villano et al., 2010) The underlying mechanisms of this cathodic extracellular
electron uptake remain less understood than its anodic counterpart, but also include
mediated EET via small molecules (e.g. H2) and direct EET via porin-cytochrome
complexes analogous to those described in Shewanella. (Lohner et al., 2014; Rowe et al.,
2019) Interestingly, some bacteria appear to be capable of bi-directional EET. S.
oneidensis, for example, is capable of directing electrons from a cathode to intracellular
electron acceptors such as fumarate or O2, allowing regeneration of ATP and NADH.
(Ross et al., 2011; Rowe et al., 2018) This process appears to rely, at least partially, on
some of the same molecules that facilitate outward EET; outer membrane multiheme
cytochromes play a role in mediating inward EET into S. oneidensis, (Rowe et al., 2018)
and flavins have likewise been observed to play a role.(Okamoto, Nakamura, et al., 2014;
26
Ross et al., 2011) However, previous studies suggest that additional enigmatic
components, beyond the canonical inward EET pathway, may play a role in directing
electrons from cathodes into S. oneidensis. (Rowe, Salimijazi, et al., 2021)
Recent evidence of oxidation of Fe2+ by an outer membrane cytochrome common
to iron oxidizing bacteria, (Keffer et al., 2021) and the recent report that redox cycling of
dissolved iron (Fe3+/Fe2+) may play a role in mediating EET from S. oneidensis to anodes,
(Oram & Jeuken, 2016) motivated us to consider whether dissolved iron can also play a
role in inward EET from cathodes to S. oneidensis. Here we present electrochemical
measurements of enhanced electron uptake by S. oneidensis in the presence of exogenously
added FeCl2 and the terminal electron acceptor fumarate. Electron uptake from the
cathode is coupled to fumarate reduction by the fumarate reductase FccA, and outer
membrane cytochromes play an important role in the inward EET pathway, as indicated
by the significantly reduced current levels in S. oneidensis mutants deficient in
cytochromes.
2.3 Materials and Methods
2.3.1 Cell cultivation
A preculture culture was cultivated from a frozen stock in 5 mL lysogeny broth (LB)
aerobically at 30°C up to an optical density (OD 600 nm) of 2.4-2.6. Defined Media (DM)
was inoculated with the washed preculture culture to a final OD of 0.05. DM contained
27
(per liter): 0.2 g MgCl2 6H2O, 1.0 g NH4Cl, 0.2 g CaCl2 2H2O, 0.9 g NaCl, 2.5 g NaHCO3,
7.2 g HEPES buffer. For this culturing stage we added 0.5 g of yeast extract, 3.316 g
Sodium lactate 60%(w/w) syrup. Trace minerals, amino acids and vitamin solution were
added to supplement the medium as described previously. (Rowe et al., 2018) The pH of
the medium was adjusted to 7.0 using NaOH. Cells were incubated at 30°C to an OD of
1-1.5. Cells were then washed 3X with fresh DM and were resuspended in 25 mL DManode (OD 0.1) with 30 mM lactate, trace minerals and amino acids for anodic cultivation
(no yeast extract or vitamins).
2.3.2 Reactor setup and electrochemical measurements
A custom-built single chamber bioreactor was used to investigate the effect of FeCl2 on
extracellular electron uptake rate by S. oneidensis MR-1. The reactor contained a graphite
working electrode (POCO AXF-5Q 0.059” x 0.225” x 0.83”) (Tri-Gemini LLC, CAT-num:
XM15839C), platinum wire as a counter electrode, and an Ag/AgCl in 1M KCl reference
electrode (CH Instruments, Inc.). The working electrode was polished using a 600-grit
sandpaper, rinsed, and sonicated twice, first in distilled de-ionized water, then in ethanol.
The working electrodes were stored in 1 M HCl until further processing. For anodic
conditions (+435 mV vs SHE) 25 mL DM contained 30 mM sodium lactate, trace minerals
and amino acids (DM-anode). For cathodic conditions (-305 mV vs SHE) 25 mL DM
contained 30 mM sodium fumarate (DM-cathode). Cyclic voltammetry (CV) was
performed by scanning the potential range of -565 mV to 635 mV (vs SHE) at a scan rate
28
of 1 mV/s. All electrochemical analysis were performed under nitrogen atmosphere, at
30°C using a CHI1000 8-channel potentiostat (CH Instruments, Inc.).
2.3.3 Biofilm formation
Biofilm formation was facilitated in anaerobic conditions by poising the working electrode
at an anodic potential (+435 mV vs SHE) with lactate serving as the electron source for
24 h. After 24 h the reactors were taken into an anaerobic chamber and electrodes were
gently rinsed and transferred into a fresh reactor containing fresh DM-cathode. The
rinsing of the electrodes ensures that only the firmly attached cells are present on the
electrode for downstream experiments. The same culturing, biofilm formation, and
electrochemical preparation was done for all S. oneidensis strains: wild-type, ∆mtrC,
∆omcA, ∆mtrC/omcA, ∆OMC and ∆fccA (Table 1).
2.3.4 FeCl2 addition
To understand the effect of soluble ferrous chloride addition on the magnitude of cathodic
electron uptake by S. oneidensis, we added anaerobic FeCl2 to a final reactor concentration
of 1 mM during chronoamperometry in the presence of fumarate. With working electrodes
poised at -305 mV (vs SHE) current was monitored for ~30 min until a steady baseline
was reached before the 1 mM FeCl2 addition. FeCl2 stock was made as previously
described. (Barco et al., 2015) To determine if the enhanced electron uptake phenotype is
dependent on free iron ions in solution we added sterile anaerobic stock of the iron chelator
29
deferoxamine (1.5 mM per 1 mM FeCl2). To investigate the FeCl2 concentration
dependence of the enhanced electron uptake in S. oneidensis, we performed
chronoamperometry and cyclic voltammetry measurements with increasing
concentrations: 0.2 mM, 0.6 mM and 1 mM FeCl2. To determine the link between Feenhanced electron uptake and fumarate reduction, we performed chronoamperometry
experiments with the working electrode poised at -305 mV vs SHE using a fumarate
reductase mutant strain (∆fccA) in the presence of fumarate. Similar conditions were
utilized for investigating the role of outer membrane and periplasmic cytochromes in Feenhanced cathodic EET (∆mtrC, ∆omcA, ∆mtrC/omcA, ∆OMC).
2.3.5 Iron chelator addition and toxicity test
During chronoamperometry 1.5 mM iron chelator deferoxamine myselate salt (DFO)
(Sigma-Aldrich Co.) was added 30-40 minutes after FeCl2 addition. Cyclic voltammetry
was performed 30 minutes after DFO addition to investigate the electrochemical
interaction between the cathode and S. oneidensis after iron chelation. Voltammetry was
performed by scanning the potential range of -565 mV to 635 mV (vs SHE) at a scan rate
of 1 mV/s. To investigate any possible toxic effects on cells due to DFO, CFU counts
were performed via serial dilutions of cells in DM-cathode media before and 30 minutes
after DFO addition. Plates were incubated in anaerobic conditions at 30°C with fumarate
serving as the electron acceptor. Colonies were quantified 24 h after incubation.
30
2.3.7 Fluorescence microscopy
After running electrochemical experiments, electrodes were processed for fluorescence
microscopy imaging. Electrodes were gently rinsed with fresh media and then fixed and
stained in 1 mL fresh DM containing 25% glutaraldehyde, and 0.25 µg/mL FM4-64FX
membrane stain solution (Thermo Fisher Scientific). Samples were stored in 4°C for ~15
h prior to imaging. Sample images were taken using the 100X objective of a Nikon Eclipse
Ti-E inverted fluorescent microscope. Images were taken from 10 fields of view per sample.
2.3.7 Tetramethylbenzidine heme stain SDS-PAGE protein gel
After anodic biofilm formation, planktonic cells were harvested and centrifuged for 10
minutes at 6,500 RCF. The pellet was resuspended and incubated in anaerobic serum
bottles containing anaerobic DM-cathode media (with fumarate) for 30 minutes before
FeCl2 addition to reach 1 mM concentration. 5 mL aliquots were taken immediately and
then again 30 minutes after addition FeCl2, centrifuged for 10 minutes at 6,500 RCF, and
resuspended in 1 mL DM-cathode media. These samples were centrifuged again for 10
minutes at 20,000 RCF and resuspended in 50 µL Laemmli sample buffer (Bio-Rad).
Subsequently, the sample was boiled for 5 minutes. 10 µL of sample was loaded for running
a 12% SDS-PAGE protein gel. The gel was rinsed with DI water for 5 minutes followed
by immersion in a solution containing 15 mL 6.3 mM TMBZ in methanol and 35 mL 0.25
31
M sodium acetate (pH 5.0). Gels were left in the dark shaking at 40 RPM for 2 h followed
by addition of 3 mL 30% hydrogen peroxide, and bands were visible within 3 minutes.
2.4 Results and Discussion
2.4.1 Iron (II) chloride enhances extracellular electron uptake by
S. oneidensis.
In order to investigate the effect of soluble iron on extracellular electron uptake by S.
oneidensis, we performed electrochemical analyses in single chamber bioelectrochemical
reactors (Figure 2.1 A) in the presence and absence of 1 mM FeCl2 with the electrode
serving as the sole electron source. In brief, following an anodic cultivation step in
anaerobic reactors (with lactate as the electron donor and the electrode as electron
acceptor), to promote biofilm formation, the electrodes were rinsed and transferred to new
reactors with fresh media for cathodic measurements. The cathodic reactors included
fumarate as the electron acceptor but no lactate (or other exogenous carbon source) to
ensure that the cathodes serve as the sole electron donor. From S. oneidensis chronoamperometry measurements, a sustained and increasing cathodic current was immediately
observed upon addition of FeCl2 into reactors with working graphite electrodes poised at
-305 mV (vs SHE) and in the presence of 30 mM fumarate as a cellular electron acceptor
(Figure 2.1 B). This cathodic current, which reached as high as -271.5 ± 41.8 µA in less
than an hour, was absent in a control including cells but no added FeCl2, and an abiotic
32
Figure 2.1. Addition of FeCl2 enhances cathodic electron uptake by Shewanella oneidensis
MR-1. (A) Schematic of the bioelectrochemical reactor. The reactor consisted of a
stoppered glass bottle with a graphite working electrode (WE), platinum counter electrode
(CE), Ag/AgCl reference electrode (RE), and N2 ports. (B) Chronoamperometry in
anaerobic cathodic conditions with 30 mM fumarate as the electron acceptor and the
working electrode poised at +305 mV (vs SHE) in the presence or absence of 1 mM FeCl2.
1 mM FeCl2 was added at the time point indicated by the black arrow. Error bars indicate
standard error of triplicate measurements. (C) Representative cyclic voltammetry mutants
(+465 to 435 mV at 1 mV/s) of S. oneidensis with 30 mM fumarate as the electron
acceptor in the presence or absence of 1 mM FeCl2.
control with FeCl2 (with current levels ~ -0.3 µA for both). It is important to note that
S. oneidensis has been previously shown to be capable of direct and flavin-mediated
electron uptake from electrodes. (Okamoto, Nakamura, et al., 2014; Ross et al., 2011;
Rowe et al., 2018) Our observation of low current densities (in the absence of exogenous
FeCl2) reflects different experimental conditions, as previous studies were performed either
in aerobic conditions (Rowe et al., 2018) or with higher cell densities and experimenting
with added flavins. (Ross et al., 2011)
33
Figure 2.2. Cyclic voltammetry scan (-465 mV to 435 mV at 1 mV/s) of abiotic control
electrodes in the presence or absence of 1 mM FeCl2.
Following this period of chronoamperometry in the presence of FeCl2, cyclic voltammetry
scan was performed to assess the electrochemical interaction between cathodes and S.
oneidensis cells. In the presence of FeCl2 and fumarate, cyclic voltammetry revealed a
clear catalytic wave with an onset potential of ~0 mV vs. SHE, a feature absent from a
control without FeCl2 (Figure 2.1 C). This onset potential is consistent with the redox
characteristics of FeCl2 in the abiotic (cell-free) control, which revealed quasi-reversible
electron transfer behavior (Figure 2.2). Fluorescence microscopy of the electrode-bound
biofilms showed (Figure 2.3) similar cell density both in the presence and absence of added
FeCl2, which reflects the anodic biofilm formation step done prior to cathodic
measurements (see Methods) and indicating that the observed enhancement in cathodic
current is the result of improved electron transfer rather than differences in biomass. The
34
Figure 2.3. Representative fluorescence microscopy images of the graphite electrode with
attached WT S. oneidensis cells stained with FM 4-64FX cell membrane stain with 1 mM
FeCl2 (A) and without 1 mM FeCl2 (B). Images were taken after 15 h of staining.
Fluorescence images shows similar cell density on the electrode surface with or without
exogenous FeCl2. This indicates that current variability between conditions is due to the
presence of soluble iron in the system.
amperometry and voltammetry measurements described above demonstrate that addition
of soluble ferrous iron significantly enhances (and can account for almost all the)
extracellular electron transfer from graphite electrodes to S. oneidensis under the
experimental conditions tested here. Although cathodic current started increasing
immediately upon iron addition and saturated in ~30 minutes, we considered the
possibility that soluble iron addition may have contributed to increased electron uptake
indirectly by impacting the expression of cytochromes. However, TMBZ heme-staining
SDS-PAGE gels of protein content, from cells harvested after the anodic cultivation step
and resuspended in the same media as the cathodic reactors, revealed no discernible
difference in cytochrome production between FeCl2 addition and 30 minutes after FeCl2
35
addition (Figure 2.4). Previous studies under comparable conditions (anaerobic with
fumarate as the electron acceptor) highlighted the role that added flavins can play as
soluble electron shuttles to enhance cathodic electron uptake. (Okamoto, Kalathil, et al.,
2014; Ross et al., 2011) Our results suggested that redox cycling of dissolved iron could
play a similar role, which next motivated us to examine the effect of added FeCl2
concentration and specific interactions with cellular redox components.
Figure 2.4. FeCl2 addition does not induce overexpression of heme-containing cytochrome
in S. oneidensis. Representative total cell protein SDS-PAGE and densitometry analysis
before and 30 minutes after FeCl2 addition. (1) Protein markers of 250, 180, 130, 95, 72,
55, 43, 34, 26, 17 & 10 kDa molecular wight; (2) total cell sample before FeCl2 addition
(3) total cell sample 30 minutes after FeCl2 addition. (A) Coomassie gel; (B) TMBZ hemestained SDS-PAGE gels and their densitometric profile.
36
2.4.2 Fe-enhanced cathodic electron uptake in S. oneidensis is
concentration dependent and due to free Fe ions.
Increases in EET mediated by soluble redox shuttles are expected to depend on the
concentration of the shuttle, as previously observed for riboflavin-enhanced EET in S.
oneidensis. (Baron et al., 2009) To that end, we investigated the concentration dependence
of Fe-enhanced electron uptake by S. oneidensis in the presence of 0.2, 0.6, and 1 mM
FeCl2 and with 30 mM fumarate serving as electron acceptor. Since Fe2+ is toxic to cells
at high concentrations, 1 mM FeCl2 was chosen as an upper bound that is tolerated by S.
oneidensis. (Bennett et al., 2015) Once again, a period of chronoamperometry (Figure
2.5A) was followed up with cyclic voltammetry (Figure 2.5B) to assess the impact of each
Figure 2.5. The FeCl2-enhanced cathodic current into S. oneidensis is concentration
dependent. (A) Chronoamperometry (+305 mV vs SHE) of S. oneidensis with increasing
concentration of FeCl2: 0 mM, 0.2 mM, 0.6 mM and 1 mM. 30 mM fumarate served as
the electron acceptor. Error bars indicate standard error of triplicate measurements. (B)
Representative cyclic voltammetry scan mutants (+465 to 435 mV at 1 mV/s) of S.
oneidensis with increasing concentration of FeCl2.
37
FeCl2 concentration on sustained inward EET and the resulting catalytic waves, in the
presence of fumarate as a cellular electron acceptor. From chronoamperometry, the
observed cathodic current was found to depend very strongly on FeCl2 concentration.
While the response to 0.2 mM FeCl2 was similar to a no-FeCl2 control, modest current
was detected at 0.6 mM FeCl2, and an order of magnitude enhancement was seen at the
1 mM FeCl2 level (Figure 2.5A). Corresponding cyclic voltammograms are shown in Figure
2.5B. While catalytic activity was not observed in the absence of FeCl2, the catalytic
waves were better defined and increased sharply in magnitude as a function of increasing
FeCl2 concentration (Figure 2.5B). The strong dependence of catalytic activity on FeCl2
concentration, and the requirement for millimolar concentration to achieve appreciable
inward EET are consistent with a redox shuttling mechanism where Fe3+/Fe2+ is
continuously oxidized/reduced by the cells and electrode respectively. The inherently
small diffusion coefficients of redox shuttles are indeed known to limit outward EET which
necessitates high shuttle concentrations to maximize the concentration gradient that
drives diffusion. (Picioreanu et al., 2007; Torres et al., 2010) It is worth noting however
that iron is an essential trace nutrient and its homeostasis is regulated by S. oneidensis
using both import (Bennett et al., 2018) and export (Bennett et al., 2015) systems which
could potentially influence the concentration available for EET mediation.
To further confirm that the Fe-dependent inward EET to S. oneidensis stems
specifically from freely diffusing Fe ions in solution, we performed experiments with added
38
free iron chelator, deferoxamine myselate salt (DFO). Following the approach suggested
by Oram and Jeuken, (Oram & Jeuken, 2016) we hypothesized that if iron acts as a free
soluble shuttle, DFO will abolish the enhanced electron uptake phenotype. Indeed, the
observed cathodic current by S. oneidensis (at -305 mV vs SHE in the presence of FeCl2)
was immediately eradicated upon the addition of 1.5 mM DFO (Figure 2.6A). Likewise,
the catalytic wave detected by cyclic voltammetry was diminished in DFO containing
reactors (Figure 2.6B). While DFO is known to chelate ferric iron specifically, it can also
influence
Figure 2.6. Fe-enhanced electron uptake by S. oneidensis MR-1 is diminished by the
addition of the iron chelator deferoxamine. (A) Representative chronoamperometry curves
of S. oneidensis on graphite electrodes poised at +305 mV (vs SHE) with 30 mM fumarate
as the electron acceptor. Arrow indicates times of FeCl2 and DFO addition. (B)
Representative cyclic voltammogram mutants (+465 to 435 mV at 1 mV/s) profiles of
cell-containing and control bare electrodes with 1 mM FeCl2, and with and without
deferoxamine.
39
the oxidation of iron prior to its complexation. (Goodwin & Whitten, 1965) Such a change
in the redox background of the experiment may underlie the appearance of a transient
anodic response after DFO addition, which occurred even in abiotic control reactors
containing 1 mM FeCl2 without cellular cathodic activity (Figure 2.6A). Regardless of
this effect, it is clear that chelating iron (specifically free extracellular iron, since DFO
cannot chelate heme iron from protein) had a significant effect in eradicating the observed
iron-mediated electron uptake. To investigate whether DFO itself has a toxic impact on
cells, which may contribute to the eradication of cellular cathodic activity, we performed
colony forming units (CFU) counts before and 30 minutes after DFO addition to cells in
the same media conditions used for cathodic measurements. This comparison revealed no
difference in cell viability as a result of DFO exposure (Figure 2.7), a finding consistent
with previous studies showing that DFO exhibits low membrane permeability, is produced
by certain bacterial species for iron acquisition, and can alleviate the toxic effects of iron
overload. (G. C. F. Chan et al., 2009; Liu et al., 2022) The DFO addition experiments
provide further evidence that the enhanced inward EET into S. oneidensis is due
specifically to freely diffusing iron ions in solution.
It is interesting to consider the possible contribution of soluble iron in mediating
extracellular electron uptake, in light of previous reports that flavins may also function as
mediators for extracellular electron uptake by S. oneidensis. (Ross et al., 2011) The
enhancement in cathodic current resulting from FeCl2 addition, observed in this study,
40
appears to be higher than the enhancement previously observed from flavin addition,
(Ross et al., 2011) although under different experimental conditions and with much higher
concentrations of added Fe (1mM) relative to flavins (1 µM). Also interesting is the
situation in a natural environment where both flavins and soluble Fe may be present and
contributing to inward or outward EET. In certain environments, such as anoxic marine
sediments, flavins were detected at low nM concentrations and showed an increasing trend
with depth; this trend was inversely correlated with dissolved Fe, which was detected in
the 20-200 µM range. (Monteverde et al., 2018) The relative contributions of each
mediator to EET would ultimately depend on their respective concentrations in specific
environments, possible redox interaction, (Zhang et al., 2020) and the specific redox
reactions occurring in that environment. Outside the natural environment, we suggest
that exogenously added soluble Fe may serve as an inexpensive mediator to enhance EET
in microbial electrochemical technologies.
Figure 2.7. S. oneidensis MR-1 is not sensitive to DFO under the experimental conditions.
CFU counts via serial dilution before and after addition of 1.5 mM DFO.
41
2.4.3 Fe-mediated electron uptake in S. oneidensis is linked to
fumarate reduction
Our electrochemical experiments include fumarate as a terminal electron acceptor for S.
oneidensis. Under anaerobic conditions, the periplasmic flavocytochrome FccA serves as
the only fumarate reductase in S. oneidensis, (Maier et al., 2003) catalyzing the reduction
of fumarate to succinate. Since previous studies of extracellular electron uptake by S.
oneidensis confirmed that this process can be coupled to fumarate reduction, we tested
whether FccA is required for the Fe-mediated electron uptake mechanism reported here.
In contrast to the significant cathodic current observed with wild-type S. oneidensis, no
current was detected from a ∆fccA mutant (Figure 2.8A) in chronoamperometry under
Figure 2.8. Fe-enhanced electron uptake in S. oneidensis MR-1 is linked to fumarate
reduction. (A) Chronoamperometry measurement at +305 mV (vs SHE) of S. oneidensis
MR-1 WT and fumarate reductase mutant (∆fccA) on graphite electrode surface in the
presence of 1 mM FeCl2 and 30 mM fumarate as an electron acceptor. Error bars indicate
standard error of triplicate measurements. (B) Representative cyclic voltammogram
mutants (+465 to 435 mV at 1 mV/s) of S. oneidensis MR-1 and ∆fccA on graphite
electrode with 30 mM fumarate.
42
similar experimental conditions (1 mM FeCl2, 30 mM fumarate, and the working electrode
poised at -305 mV vs. SHE). Likewise, the cyclic voltammetry measurements of ∆fccA
were similar to the abiotic (cell-free) control, rather than the catalytic wave exhibited by
wild-type S. oneidensis (Figure 2.8B). In the absence of fumarate with 1 mM FeCl2, the
abiotic control and cell containing reactors also demonstrated similar electrochemical
signatures (Figure 2.9). This requirement for the fumarate reductase FccA and the
presence of fumarate confirm that the electrode-sourced and Fe-mediated inward EET
can enter the cellular periplasmic space to drive reduction of fumarate.
Table 2.1. Strains used to study the role of S. oneidensis outer membrane cytochromes
in FeCl2-enhanced extracellular electron uptake.
Strain Characteristics Source
MR-1 S. oneidensis wild-type
JG731 S. oneidensis MR-1, ΔmtrC (Coursolle &
Gralnick, 2010)
JG719 S. oneidensis MR-1, ΔomcA (Coursolle &
Gralnick, 2010)
JG749 S. oneidensis MR-1, ΔmtrC/omcA (Coursolle &
Gralnick, 2010)
JG686 S. oneidensis MR-1, ΔfccA (Ross et al.,
2011)
JG1486 S. oneidensis MR-1, ΔomcA/ΔmtrA/ΔmtrF/
ΔdmsE/ΔSO4360/ΔcctA/ΔrecA (ΔOMC)
(Coursolle &
Gralnick, 2012)
43
Figure 2.9. Cyclic voltammetry scans (+465 to 435 mV at 1 mV/s) of S. oneidensis MR1 and sterile reactors with 1 mM FeCl2 in the absence of electron acceptor fumarate.
2.4.4 Fe-mediated electron uptake is dependent on outer membrane
and periplasmic cytochromes
Previous studies of cathodic electron uptake by S. oneidensis implicated the reversibility
of the same Mtr pathway responsible for catalyzing outward EET to minerals and anodes.
(Ross et al., 2011; Rowe et al., 2018) To examine whether the Fe-mediated process
interacts with the outer membrane cytochromes, which in turn pass electrons to the
periplasmic cytochromes and ultimately to FccA, we performed electrochemical
experiments comparing Fe-mediated electron uptake by wild-type S. oneidensis and
mutant strains lacking key Mtr pathway components (Table 1): ΔmtrC, ΔomcA,
ΔmtrC/omcA, and ΔOMC (ΔomcA/ΔmtrA/ΔmtrF/ΔdmsE/ΔSO4360/ΔcctA/ΔrecA).
Figure 10A displays the Fe-mediated inward EET currents from the tested strains with
the working electrode poised at -305 mV vs. SHE. The mutant strains lacking either of
44
Figure 2.10. Fe-enhanced cathodic electron uptake is catalyzed by outer membrane
cytochromes in S. oneidensis MR-1. (A) Chronoamperometry experiments (-305 mV vs
SHE) of S. oneidensis, mtrC mutant (ΔmtrC), omcA mutant (ΔomcA), mtrC and omcA
double mutant (ΔmtrC/omcA), and outer membrane cytochrome (OMC) mutant
(ΔomcA/ΔmtrA/ΔmtrF/ΔdmsE/ΔSO4360/ΔcctA/ΔrecA) with 30 mM fumarate
serving as an electron acceptor. Addition of 1 mM FeCl2 is indicated by the arrowhead.
Error bars indicate standard error of triplicate measurements. (B) Representative cyclic
voltammograms of S. oneidensis WT and outer membrane cytochrome mutants (+465 to
435 mV at 1 mV/s) under nitrogen atmosphere.
the outer membrane decaheme cytochromes MtrC and OmcA (ΔmtrC and ΔomcA)
demonstrated cathodic current levels 37% and 42% lower than wild-type S. oneidensis,
respectively. A double deletion strain lacking both MtrC and OmcA (ΔmtrC/omcA)
exhibited an even lower cathodic current – about 73% less than wild-type levels. Electron
uptake was severely diminished in a strain lacking genes encoding eight functional
periplasmic and outer membrane cytochromes (ΔOMC), with current levels about 96%
lower than wild-type S. oneidensis.
45
Cyclic voltammetry (Figure 2.10B) echoed the chronoamperometry measurements,
showing the highest catalytic activity in wild-type S. oneidensis and proportional
reduction in the magnitude of the catalytic wave resulting from cytochrome deletions.
Additionally, fluorescence microscopy (Figure 2.11) of the electrode-bound biofilms did
not reveal significant differences between the strains, confirming the differences in cathodic
electron uptake cannot be simply attributed to different cellular coverage. Taken
collectively, these results indicate that MtrC and OmcA are the primary cell surface
conduits through which electrons are passed from the dissolved Fe into the cells, and that
the reversible Mtr pathway is the primary route for Fe-mediated inward EET.
It is interesting to consider the energetics of the interaction between soluble iron
and the cell surface multiheme cytochromes. Taking MtrC as the representative cell
surface conduit, previous protein film voltammetry (PFV) measurements show some
variation depending on experimental conditions, (Firer-Sherwood et al., 2008) but at pH
7 it can be reduced over a potential window from +100 mV to -400 mV vs SHE,
(Hartshorne et al., 2007) with this broad range reflecting the presence of the 10 heme
centers with overlapping potentials. Moreover, calculations of the individual heme
reduction potentials show that this window can shift depending on the overall reduction
state of the other hemes in the protein. (Barrozo et al., 2018; Jiang et al., 2019) The
reduction potential of the iron mediator depends on its interactions with other ligands
46
Figure 2.11. Representative fluorescence microscopy images of the graphite electrode with
attached S. oneidensis WT (A) and mutant cells stained with FM 4-64FX cell membrane
stain after electrochemical cultivation and cyclic voltammetry with added FeCl2. Images
were taken after 15 h of staining. Fluorescence images shows similar cell density on the
electrode surface with exogenous FeCl2. This indicates that current variability between
conditions is due to the availability of heme containing cytochromes that shuttles electrons
into the cell.
47
(e.g. fumarate in our experiments) and the particular electrode material, (Oram & Jeuken,
2016) but our measurements show a catalytic onset potential for the Fe-mediated
mechanism of ~ 0 mV vs. SHE (Figure 2.1). This value is at the higher end of the potential
window from PFV measurements of purified MtrC (and OmcA), suggesting a weak driving
force for electron transfer from Fe2+ to cytochromes. However, it is also important to
consider that the reduction potentials measured with direct live cell voltammetry of
Shewanella are shifted higher than those obtained from purified cytochromes, (Nakamura
et al., 2009; Xu et al., 2016) which would result in a higher driving force for Fe-mediated
electron injection into cells.
2.5 Conclusions
We have demonstrated that soluble iron (exogenously added as FeCl2) significantly
enhances extracellular electron uptake from graphite electrodes by the model EET
organism Shewanella oneidensis MR-1. Experiments with the iron chelator deferoxamine
indicate that the enhancement is due to freely diffusing ions acting as redox shuttles,
where Fe3+/Fe2+ is continuously oxidized/reduced by the cells/electrodes. While
extracellular electron uptake was previously demonstrated in S. oneidensis, and a role for
soluble iron in mediating anodic EET was previously suggested, our study highlights how
soluble iron can also facilitate cathodic EET into S. oneidensis. In addition, we
demonstrated that the Fe-mediated inward EET is largely routed into the cells through
the same cytochrome network responsible for outward EET and can be coupled to
48
fumarate reduction by S. oneidensis through the activity of the periplasmic fumarate
reductase. These findings have implications for microbial electrochemical technologies,
including electrosynthesis where electrode-sourced currents drive microbial biosynthetic
pathways for production of high value chemicals and fuels.
49
Chapter 3: Investigation on
Microbe Mineral interaction using
electrochemical cultivation at the former
Homestake Gold mine in South Dakota
This chapter has been adapted from (Rowe, Abuyen, et al., 2021):
Rowe, A. R., Abuyen, K., Lam, B. R., Kruger, B., Casar, C. P., Osburn, M. R., ElNaggar, M. Y., & Amend, J. P. (2021). Electrochemical evidence for in situ microbial
activity at the Deep Mine Microbial Observatory (DeMMO), South Dakota, USA.
Geobiology, 19(2), 173–188. DOI: 10.1111/GBI.12420 URL:
https://doi.org/10.1111/GBI.12420
50
3.1 Abstract
Microbes are the dominant life forms in the subsurface. To investigate this environment
and the potential for deep ground water microbial community to utilize extracellular
electron transfer for respiration, we performed a long-term electrochemical cultivation at
the Deep Mine Microbial Observatory (DeMMO) specifically DeMMO 4, located in the
Sanford Underground Research Facility (SURF) in Lead, South Dakota (USA). Graphite
and Indium Tin Oxide (ITO) electrodes, maintained at a potential of -200 mV versus the
Standard Hydrogen Electrode (SHE), were subjected to long-term incubation alongside
open circuit controls and various minerals within a parallel flow reactor. We monitored
the current via chronoamperometry measurements and performed cyclic voltammetry
analysis on a weekly basis to have a better of understanding of the changes that’s occurring
in the microbial community. Here we found both microbial anodic and cathodic activity
demonstrated in both chronoamperometry and cyclic voltammetry measurements.
Additionally, catalytic activities in the cyclic voltammetry measurements were observed
with features that are very similar to pure isolates. While the microbial composition was
different between fracture fluids and surfaced attached samples, minimal difference was
observed between electrodes and minerals incubated at the site. This suggests that in this
environment, the ability to attach to surfaces is a stronger driver in microbial composition
rather than type or redox activity of the mineral. In this study we demonstrated that
51
electrochemical techniques can be implemented in situ and can provide insights into
microbial activities in the subsurface environments.
3.2 Introduction
In terrestrial deep subsurface environments, microbes represent a large portion of the
Earths biomass. (Casar et al., 2020; Jangir et al., 2019; Osburn et al., 2020) They play a
critical role in the biogeochemical cycles of carbon, nitrogen, sulfur, and iron. (Bradley et
al., 2018; McKay et al., 2016; Osburn et al., 2014) Ultimately, in these energy limiting
conditions, microbe-mineral interactions can shape microbial community interactions and
likewise influence the geochemistry of the Earth’s surface. (Emerson, 2016; Osburn et al.,
2014; Purkamo et al., 2017) However, monitoring microbial activity in real-time in situ
can be challenging. Majority of investigations in the terrestrial environments (and marine)
are based on sampling, sequencing, and genomic analyses, where microbial functions and
their environment-specific responses are currently being understood through functional
inferences from metagenomes or other indirect measurements. (Headd & Engel, 2013;
Suzuki et al., 2018) However, certain species will only express sets of functions that drive
their own fitness, via synergistic or antagonistic interactions with other species. This, in
turn, creates a disconnect between the genomic potential and the phenotype of the species
during growth in a community, making genomic and metagenomic analyses of
microbiomes insufficient to gain knowledge of the ecological roles and functions of these
communities. Though this approach has advanced our understanding with respect to the
52
diversity of the of microbes and their composition in the environment there are still major
gaps missing, particularly the link between genomic information and microbial activity in
environmental settings. Studies investigating the relationship between microbiology and
geochemical observations has provided great insight into select microbial processes,
however, these studies were done on soluble substrates such as O2, NO3
-
, SO4
2- and S2-
.
On the contrary, our understanding of metabolic interactions with solid phase substrates
in the environment is relatively less understood.
In marine and subsurface environments, mineral-based microbial metabolisms are
particularly understudied. In these environments, another challenge surfaces, despite the
limited energy source microbes are able to survive. Microbe-mineral interactions are likely
playing a critical role in microbial persistence, however, this can be difficult to quantify.
Additionally, making the connection between particular microbial behaviors that facilitate
biomass production has proven difficult, as there is a severe shortage of field-derived and
in situ measurements of activity associated with the low energy sources in subsurface
environments. Moreover, the limited access to subsurface sites has contributed to the
limitation of in situ analysis of microbial activity in these environments.
In 2016, located within the former Homestake Gold Mine, the Deep Mine Microbial
Observatory (DeMMO) was recently established at the Sanford Underground Research
Facility (SURF), Lead, SD, USA as a natural laboratory. DeMMO comprises six boreholes
at approximately 1,500 meters deep and with varying lithologies. The aqueous profiles of
53
each site is distinguishable, indicating variations in depth, mineral composition of host
rock, and proximity to mine operations. A combination of studies, including geochemical
analysis, community analysis and metagenomic sequencing provide significant evidence
for chemolithotrophic metabolisms in addition to nitrogen and sulfur cycling. (Casar et
al., 2020; Momper et al., 2017b; M. R. Osburn et al., 2014)
Microbe-mineral interaction can be mimicked using poised electrodes to drive
microbial metabolism using bioelectrochemical systems (BES). BES have become an
important tool in understanding cellular electron transfer by microorganisms in the
environment allowing researchers to control activity and redox potential in the
environment where the bacteria resides. (Jangir et al., 2016, 2019; Lam et al., 2019; Rowe
et al., 2014) Moreover, when performed in situ, this method opens doors as it allows
researchers to maintain the natural environmental conditions of the microbial community,
including temperature, pH, nutrient availability and more importantly, interaction with
other organisms. Additionally, observations over extended periods provide insight into
the temporal dynamics and adaptation to environmental fluctuations, making real-time
observations and monitoring of the microbial community advantageous. That said, from
an environmental microbiology perspective, electrocultivation techniques is well positioned
to fill some of the gaps that remain when studying microbial communities in situ.
The purpose of this study was to demonstrate and quantify microbial
electrochemical activity in situ to understand the microbe-mineral interaction at this site.
54
Figure 3.1. Graphical representation of the geographical location of DeMMO 4. Crosssectional view of the Deep Mine Microbial Observatory (DeMMO). Black lines are tunnels
and shafts in the mine. Pink circles represent locations of six DeMMO sites, and larger
circle represent location of DeMMO 4. Illustration by Caitlin Casar (Casar et al., 2020).
Moreover, we wanted to determine the microbial community responsible for such activity.
One of the six spatially distributed sites with naturally draining fracture fluids was used
as a sample source, DeMMO 4. (Osburn et al., 2019) DeMMO 4, located 4,100 ft level,
was chosen for its predicted capability to support sulfate reduction and sulfide oxidation
metabolisms. This project led to two incubations, a preliminary incubation in May 2017,
followed by another incubation in October 2018. We combine 16S rRNA analysis and
electrochemical cultivation to understand some of the mechanisms that play an important
role in the structure of the microbial community in the deep subsurface. Both anodic and
cathodic activities were observed at DeMMO 4. ITO electrodes demonstrated higher
anodic activity, while graphite electrodes were more cathodic. This study also revealed
little variability in microbial composition between electrodes, and minerals incubated at
55
the site. Taken together, these observations suggest that community composition is not
driven by the poised electrodes or minerals available but rather by the ability of the
microbes to attach to surfaces.
3.3 Materials and Methods
3.3.1 Site description
SURF located within the former Homestake Gold mine founded in 1877 and was taken
out of operation in 2002, is now one of the worlds premier underground research laboratory
managed by the South Dakota Science and Technology Authority as an underground research facility. In 2016, scientists completed the installation of DeMMO, a set of six legacy
boreholes at different depths (800′–4,850′ or 240–1,455 m) that naturally drain fracture
fluids isolated from mine atmosphere through permanent packers (Osburn et al., 2019).
In this study, electrochemical cultivation was done at the 4,100’ (1,250 m) level, DeMMO
4 (Figure 3.1). Installation of electrical power and internet at DeMMO 4 permitted the
remote operation and monitoring of electrochemical instrumentation. A pilot study was
performed February-May 2017, followed by a more robust experimental installation in
August-October 2017.
3.3.2 Geochemical analysis
Geochemical analyses were performed at DeMMO 4 on borehole/ packer derived fluids
both pre- and post-electrochemical incubations (Table 1), as previously described (Osburn
56
et al., 2019). Conductivity, pH, total dissolved solids (TDS), and oxidation–reduction
potential (ORP) were measured with a portable Myron Ultrameter II (Myron L Company)
during each sampling trip to DeMMO 4. Filtrate (<0.2 µm) was collected into acid washed
and combusted (4 hr at 450°C) amber glass vials and acidified to pH 2 with 6N HCl and
then stored at 4°C until dissolved organic carbon (DOC) analysis. Analysis of DOC was
performed by Anatek Labs, using method SM 5310B. Redox sensitive ions were measured
on site using a portable Hach DR1900 Spectrophotometer (Hach Company) and associated
Table 3.1 Geochemical measurements from DeMMO 4 at SURF, before and after
electrochemical cultivation
Feb. 15,
2017
M ay 9,
2017
Aug.
31,
2017
Oct.
16,
2017
Fluid Flow Rate (mL/min) 850 580 700 350
Temp (°C) 22.6 22.1 22.5 22.6
pH 7.28 8.22 8.36 8.38
ORP (mV) -200 -172 -255 -154
Cond (µS/cm) 1,767 1,777 1,777 1,773
TDS 1,290 1,296 1,300 1,296
Nitrate (mg/mL) 0.9 1.1 0.9 0.8
Ammonia (mg/mL) 1.24 1 1.24 1.24
Ferrous Fe (mg/mL) 0 0 0.01 0
Sulfide (mg/mL) 864 678 916 830
Sulfate (mg/mL) 289 294 284 286
Dissolved Oxygen (mg/mL) 0.027 0.027 0.031 0.017
DOC (mg/mL) 0.168 0.2 0.232 0.117
57
Figure 3.2. Schematic diagram of parallel flow electrochemical reactor. Electrodes and minerals
were placed in glass chambers filled with water from the manifold. Chambers were then submerged
in water from the borehole to prevent direct intrusion of ambient air, while still allowing fluid
flow. Passive fluid flow from DeMMO 4 manifold was delivered to glass chambers, and fluid
escaped these chambers through a 7/8-inch diameter hole in the rubber stopper (not illustrated).
The reference electrode (Ag/AgCl in 1 M KCl) and counter electrodes (titanium wire) were
submerged in glass reactor effluent. A separate counter electrode was used for each channel, though
only one is illustrated for clarity. Duplicate ITO and graphite electrodes were poised at ~-200 mV
versus SHE, along with open circuit (OC) controls.
reagent kits. Hydrogen sulfide was measured by the methylene blue colorimetric method
(Hach method 8131). Ferrous iron concentrations were measured using the 1,10
phenanthroline method (Hach method 8146). Nitrate concentrations were measured using
the NitraVer5 cadmium reduction method (Hach method 8039), and ammonia
concentrations were measured using the salicylate colorimetric method (Hach method
8155). Dissolved oxygen concentrations were also measured with low and high range
ampules (Indigo Carmine Method, Hach methods 8316 and 8166, respectively).
58
3.3.3 Mineral and electrochemical incubation
In this study, we designed a bioreactor for electrode incubations that was continuously
fed fluids from the DeMMO 4 borehole to assess in situ activity in these fluids. The reactor
was designed to prevent the intrusion of other micro-organisms and exclude atmospheric
oxygen intrusion to limit alteration of fluid chemistry as much as possible. Additionally,
cross contamination of biofilms from different incubations was minimized by splitting
flows into distinct chambers. Previous electrochemical incubations (December 2014–May
2015) investigated a range of cathodic to anodic redox potentials (530 mV to −190 mV
vs. SHE) and surprisingly the overarching microbial community of these incubations did
not vary dramatically—potentially due to a lack of barriers (between electrodes) for
microbial migration (Jangir et al., 2019). To prevent transfer or cross contamination of
the enriched microbial community members between electrodes and to stimulate
enrichment of a specific mineral or electrode, we designed a reactor that split fluid flows
from DeMMO 4 into eight different chambers (Figure 3.2). We utilized materials with
the lowest oxygen permeability available (PTFE, Viton) and glass tubing with butyl
rubber stoppers and PTFE or PVC fittings. The DeMMO 4 manifold (borehole packer)
was connected to the 8-chamber reactor via Viton tubing and PTFE fittings and
connectors. Each chamber was composed of a ~13.2 cm glass cylinder containing a ~48
ml volume, enclosed on each end with a hole-punched butyl rubber stopper fixed with
silicone caulk. For the first incubation (February–May 2017), PTFE tubing was used to
59
connect Viton tubing to the manifold. This tubing was replaced with a PVC ball valve
for the second incubation allowing us to modulate flows into the reactor during
installation. Chambers were autoclaved in individual bags and transported to SURF.
Viton tubing and valves were pasteurized at 80°C for one hour in a strong base (0.5 M
NaOH), as this tubing is non-autoclavable. A poised potential electrode, open circuit
control or mineral was added to each chamber (overview Figure 3.2). There was no overlap
in materials or conditions per chamber for incubations. DeMMO 4 fracture fluid was
directed to flow into each chamber via Viton tubing with Y-shaped connectors. Each
connector was leveled and monitored for even distribution of fluid flow. Gravitational flow
of fluids into and through these chambers was continuous. Equal flow rates to each tube
were determined at the beginning of each experiment by quantifying the fluid output in
each chamber over time and modulating flow rates until equivalent flow rates were
achieved. Fluids escaped from the bottom of each chamber through a size 9 rubber stopper
with a 7/8-inch diameter hole to prevent limitation of fluid flows (inflow tubing 11/18-
inch inner diameter). All chambers were submerged in a secondary container that
prevented air intrusion into the chambers (overview in Figure 3.2). However, gas exsolving
from the DeMMO 4 fluids could enter the system, resulting in headspace accumulation in
each of the chambers. Though variability of the gas volumes in each reactor was noted,
no reactor accumulated gas below the secondary container fluid interface, and the
incubated minerals and electrodes remained submerged in fluids. Gases exsolving from
60
fluid at this site include ethane, methane, carbon dioxide, carbon monoxide, hydrogen,
and helium. (Osburn et al., 2019)
In each experimental replicate, four chambers were devoted to electrochemical
experiments. Indium Tin Oxide (ITO) electrodes (7.62 × 2.54 cm, 5–15 Ω per cm2 plated
glass, Delta Technologies [product number CB-50IN-1507]) were cleaned using ethanol
and connected to a titanium wire at a single point using silver paint and insulated using
marine epoxy. Graphite electrodes (2.54 × 2.54 × 0.254 cm, McMaster Carr [product
number 9121K67]) were sonicated in 99% EtOH for 20 min. For connection, a titanium
wire was looped around through a 21 G hole. Electrodes were then autoclaved in autoclave
bags. Coiled titanium wire counter electrodes (~1 m) were polished using 800 grit
sandpaper, rinsed with acetone, and flamed to remove organic material prior to
autoclaving. Silver/silver chloride (Ag/AgCl) reference electrodes (1 M KCl, CH
instruments [CHI111]) were used as reference electrodes. Due to design constraints the
distance between the working electrode and reference was minimized but not uniform,
which may result in subtle shifts in redox potentials at the working electrode due to iR
drop. However, we observed no evidence for this effect in CV and open circuit analyses,
and the conductivity of the fluids should minimize this effect. The other four chambers
were incubated with open circuit controls and various mineral treatments—specifically,
elemental sulfur, pyrite, and quartz were used as these have been previously associated
with geology of the formations present in the Homestake mine. (Caddey et al., 1991)
61
Pyrite and sulfur minerals were sliced into coupons using an IsoMET low speed
saw (Buehler) to obtain a flat surface. These mineral coupons were rinsed with ethanol
then dried under UV while turning periodically for sterilization. Quartz minerals were
baked at 400°C overnight and then autoclaved. Though previously measured fluid flows
had remained relatively consistent over our periodic sampling of DeMMO 4, we noted in
our May sampling (at the end of the Feb.-May incubation) the borehole packer at DeMMO
4 was leaking fluid around the edges. Consequently, the amount flowing through the
packer was significantly reduced (from 850 to 580 ml/min). While we do not know the
specific timing of this event, our electrochemical data suggest this was likely in the last
few weeks of our incubation (described in Results). This also resulted in a change in
hydraulic residence time (flow rate/reactor volume) from 2.2 per min to 1.5 per min. The
bushing on the packer was replaced prior to the Aug.-Oct. incubation which increased the
fluid flow back to 700 ml/min. However, over the course of the second incubation, the
fluid flow rate also decreased (to 350 ml/min). This decrease could not be linked to packer
leakage, though a change in hydraulic residence time from 1.8 to 0.9 was noted.
Nonetheless, the fluid flow rates did not appear to dramatically change the native chemical
composition of most of the anions observed from the borehole fluids, though we were not
able to monitor changes in geochemistry to the internal reactor environments. Differences
in the hydraulic residence time of these fluids could have altered the effective
concentrations in terms of nutrient available to microbial populations and resulted in
variation in the activity observed over time across treatments. Though we could not
62
directly monitor fluid flows over the course of these experiments, it is likely that variability
in fluid flows may help explain some of the fluctuations and heterogeneity observed in the
electrochemical data as described below.
3.3.4 Electrochemical data analysis
Prior to running long-term amperometry experiments in this reactor setup, we used linear
sweep voltammetry to assess the potential for abiotic electrode interactions on the preincubation electrodes in DeMMO4 fracture fluids (Figure 3.3). For example, depending
on the electrode redox potential and the geochemical conditions, redox active ions such
as iron or sulfide can interact with electrodes. Reduction current was observed at redox
potentials upward of −100 mV versus SHE on both the graphite and ITO electrodes
(Figure 3.3). This is consistent with the redox potential where we expected sulfide to be
electrochemically oxidized to sulfur on the electrode based on energetics and previous
observations. (Rowe et al., 2015) Given that little to no electrochemical activity was
observed below −200 mV versus SHE, this potential was utilized to minimize the influence
of abiotic signal in the long-term measurements. Notably, this also matched the prevailing
redox conditions measured at the site (Table 3.1). In the first incubation (February–May
2017), four potentiostat channels were used to test two graphite and two ITO electrodes
poised at −200 mV versus SHE.
63
Figure 3.3. Pre-incubation linear sweep voltammograms (LSV) of filtered DeMMO4
fracture fluids using indium tin oxide (ITO). Scan rate at 0.1 mV/s within the range of
-600 mV to +500 mV (SHE).
Replicate open circuit controls were performed for each electrode type. In the
second incubation (August–October 2017), an improved reactor design allowed better flow
regulation and the addition of more chambers. In this case, eight potentiostat channels
(controlling four ITO and four graphite electrodes) poised the electrodes at −200 mV vs
SHE. Both incubations were performed using a CHI1000 eight channel potentiostat (CH
Instruments, Inc.). For each incubation, Ag/AgCl (1 M KCl) reference electrode, and
titanium wire (~1 m) counter electrodes were used (~12 cm2 surface area). Voltage of the
reference electrode was quantified before and after experiments to a standard reference
and each electrode was within 15 mV of the standard (−222 mV vs. SHE), except for our
first experiment where a larger deviation was noted approximately two months of
incubation. The references for all channels in the first experiment were changed at this
time. Current measurements at −200 mV (chronoamperometry) were performed at 1-min
64
sampling intervals for each electrode over the 6–9-week incubations. Prechronoamperometry, Linear Sweep Voltammetry (LSV [0.1 mV/s scan rate from −600 to
+500 mV]) and Cyclic Voltammetry (CV [0.1 and 1 mV/s scan rate, May and Oct.,
respectively, over a −600 to +500 mV range]) were performed for both incubations. To
obtain better microbial activity resolution, weekly CV scans were run during the second
incubation along with a final post-experiment CV. CV was run over a range of −600 mV
to +500 mV versus SHE at 1 mV/s unless indicated otherwise.
3.3.5 Sample collection and DNA extraction
After long-term incubations, we collected biofilm samples that formed on the electrodes,
open circuit controls, and minerals, in addition to filtered fluid samples from the DeMMO
4 manifold. After deconstruction of the reactor, biofilm and filter samples were transferred
to sterile whirl pack bags for fixation in formalin (~10 min). To prevent any DNA
degradation from fixation, samples were immediately stored on ice or in a 4°C refrigerator
until the samples were shipped on dry ice to the laboratory for processing. (Koshiba et
al., 1993) There, the biofilm material was removed by scraping the surface of the substrate
using a sterile razor blade. Samples were then collected in microfuge tubes, centrifuged at
13,000 g for 10 min, and processed for DNA extraction using the MoBio PowerSoil DNA
isolation kit (MoBio). DeMMO 4 water samples were also taken by filtering through a
0.2-µm Sterivex filters (Milli- pore, Sigma) onsite, and filters were immediately placed on
dry ice for transport to the laboratory. As all liquid could be removed effectively from the
65
filters, filter samples were not formalin fixed before freezing and extraction. Filters were
stored at −20°C until further processing. Water samples from February, May, and August
2017 were extracted directly from the intact Sterivex filters using MoBio PowerWater
Sterivex DNA isolation kit following the manufacturer's protocol. The October 2017 water
sample was aseptically cut into pieces and extracted using the MoBio DNA PowerSoil
DNA isolation kit (MoBio). Blank extractions were performed as a negative control for
potential contamination. Quality and quantity of the extracted DNA were determined via
absorbance spectroscopic method using a Nanodrop (Thermo Fisher Scientific). Extracted
DNA was concentrated for sequencing using Zymo research purification kit (Zymo
Research), prior to submission for 16S rRNA-tagged gene amplicon sequencing.
3.3.6 Microbial community analysis
DNA extracts from electrodes, minerals, planktonic biomass, and the October 2017 water
sample were sequenced using 16S rRNA-tagged amplicon Illumina sequencing by MRDNA (Molecular Research LP). The water samples collected from February, May, and
August 2017 were sequenced at Argonne National Laboratory as described previously.
(Casar et al., 2020) Using the 16S rRNA V4 variable region PCR primers 515/806
(Caporaso et al., 2011; Soergel et al., 2012), and approximately 20 ng of environmental
genomic DNA, a 28 cycle PCR was performed at the following conditions: 94°C for 3 min
hotstart, followed by 28 cycles at 94°C for 30 s denaturation step, annealing step at 53°C
for 40 s and elongation step at 72°C for 1 min, with a final elongation step at 72°C for 5
66
min. Samples were pooled in equal proportions and purified using Ampure XP beads
(Agencourt Bioscience Corporation). Sequencing was performed using MISEQ (Illumina)
chemistry and instrumentation following manufacturer's guidelines. The Quantitative
Insights Into Microbial Ecology (QIIME v. 1.9.1.) bioinformatics pipeline was used for
barcode removal, quality filtering, and chimera detection and removal. (Bokulich et al.,
2013; Caporaso et al., 2010; Edgar et al., 2011) De novo operational taxonomic unit (OTU)
picking was utilized for taxonomic assignment (based on 97% similarity), and sequence
alignment. (Caporaso et al., 2010) OTUs associated with negative control samples (blank
extraction) (DNA extraction kit processing) were also filtered and removed from
experimental samples (in total, 985 OTUs out of nearly 50,000 OTUs were removed). The
R package Phyloseq was utilized for downstream analyses of 16S rRNA sequencing data
including principal coordinates analysis (PCoA) (McMurdie & Holmes, 2013) and the
DESeq2 analysis for differential enrichment. (Love et al., 2014; McMurdie & Holmes,
2014) To avoid artifacts due to differences in sequencing depth between samples, sequence
data were rarefied to the sample with the lowest sequence yield (n = 6,438). We used
rarefied data and normalized to equivalent number of sequences for each sample and
report relative abundance calculations and principal coordinate analyses from these data.
Non-normalized OTUs observed in this work are provided in Table S1. The raw sequences
have been uploaded to the NCBI Sequence Read Archive (SRA) database (Bioproject ID:
PRJNA577728).
67
3.4 Results and Discussion
3.4.1 Evidence of both oxidizing and reducing microbes
(1st incubation)
To investigate the activity of microbe–mineral redox interactions at DeMMO 4, a series
of mineral and electrochemical incubations was performed, using poised potential
electrodes (-200 mV vs. SHE). The pilot incubation of 9 weeks from February through
May 2017 showed evidence of electrochemically active microorganisms present at DeMMO
4. (Rowe, Abuyen, et al., 2021) In that work, predominantly negative currents were
measured, which indicates the occurrence of electron uptake. In a similar study in marine
sediments, negative currents have been observed at a similar potential (Rowe et al., 2014)
and at lower potentials. (Jangir et al., 2019; Mohamed et al., 2019) In both of these
studies, cathodic (negative) current increased over time. However, as chronoamperometry
data can be the result of abiotic reactions, biotic reactions, or both, electrochemically
active microbes were ultimately isolated and characterized from these incubations to
support biological activity.
In addition to the chronoamperometry measurements, the preliminary work also
demonstrated evidence for microbial catalysis at the electrode surface via cyclic
voltammetry measurements. (Rowe, Abuyen, et al., 2021) A distinct difference was
observed between electrodes. The ITO electrodes demonstrated an anodic catalytic wave
feature which indicates the coupling of electron donor oxidation to anode reduction. This
68
catalytic signature is similar to that of pure metal reducing cultures such as Geobacter
sulfurreducens, (Marsili et al., 2008, 2010) and is distinct from what would be expected if
a redox active diffusible molecule, such as iron or sulfide, was reacting with the electrode.
This suggests that there is microbial solid phase mineral reductive metabolisms on the
electrode surface. While ITO electrode 1(ITO-1) showed anodic catalytic activity, graphite
electrode 1 (GRA-1) demonstrated cathodic catalytic activity, specifically coupling
cathode oxidation to the reduction of an electron acceptor in the reactor. (Rowe, Abuyen,
et al., 2021) This is not surprising given the predominant cathodic current observed in
the chronoamperometry measurements. Though the redox potential of the electron uptake
may be different, this catalytic feature is similar to those observed in cathode-oxidizing
pure cultures. (Karbelkar et al., 2019; Rowe et al., 2014) For example, previous work
investigating electron uptake from marine sediments demonstrated a near 500 mV range
in redox potentials for electron uptake features in environmental samples. (Lam et al.,
2019; Rowe et al., 2014; Yates et al., 2016) First derivative analysis of the anodic and
cathodic sweeps of CVs demonstrated mid-point potential peak ranges between −206 to
−118 mV and −201 to −146 mV for ITO-1 and GRA-1 electrodes, respectively. This
suggests that perhaps that anodic and cathodic biofilms can operate in a similar redox
potential window. Indicating the possibility of multiple electrochemical processes
operating in the fluid regime. Another possible explanation would be that there are
reversible electrochemical pathways present in this environment.
69
Towards the end of the preliminary incubation, current levels in both ITO and
Graphite reactors drifted toward more positive currents. We associated this with the
reduced fluid flow due to the DeMMO 4 packer. This ultimately led to reduced mixing in
the reactors and/or increased hydraulic residence time driving oxidant (i.e., O2, NO3- )
limitation at the electrode generating more positive currents. It should also be noted that
these amperometry data reflect the net electron flow, which are likely a cumulation of the
dominant redox process occurring at any given time. That said, changes in current likely
reflect real-time shifts in the environmental conditions as well (organic carbon abundances,
oxygen concentrations, etc.) rather than just microbial community influence. However,
we cannot rule these out as affecting current data. While the CV analysis points to
microbial activity, based on the chronoamperometry measurements there were significant
changes in the direction and magnitude of the current measured over the course of the
entire electrochemical incubation. The observed variability can possibly be explained by
spatial and temporal variability in geochemistry (abiotic reactions), microbial community
abundance/structure varying with fluid flows to the reactor and/or responding to
variation in geochemistry (biotic reactions), or both. To address some of the temporal
variability in this pilot work, and whether or not differences observed over time on these
electrodes are a result of biological activity, our second reactor installation expand our
electrode replicates, and modified the reactor for better fluid flow control.
70
Figure 3.4. Chronoamperometry profile and cyclic voltammetry plots of Indium tin oxide
1 electrode, ITO-1. Chronoamperometry profile is shown for the 45-day incubation which
were poised at ~−200 versus SHE. Black circles indicate CV runs. (A). Data was recorded
at 30 s intervals. Cyclic voltammetry plots run with a scan range of −600 mV to +300
mV versus SHE with a scan rate of 1 mV/s: day 7 (B), day 14 (C), day 21 (D), day 39
(E), and day 45 (F).
71
Figure 3.5. Chronoamperometry profile and cyclic voltammetry plots of Indium tin oxide
3 electrode, ITO-2. Chronoamperometry profile is shown for the 45-day incubation which
were poised at ~−200 versus SHE (A). Data was recorded at 30 s intervals. Cyclic
voltammetry plots run with a scan range of −600 mV to +300 mV versus SHE with a
scan rate of 1 mV/s: day 7 (B), day 14 (C), day 21 (D), day 39 (E), and day 45 (F).
72
Figure 3.6. Chronoamperometry profile and cyclic voltammetry plots of Indium tin oxide
3 electrode, ITO-3. Chronoamperometry profile is shown for the 45-day incubation which
were poised at ~−200 versus SHE (A). Data was recorded at 30 s intervals. Cyclic
voltammetry plots run with a scan range of −600 mV to +300 mV versus SHE with a
scan rate of 1 mV/s: day 7 (B), day 14 (C), day 21 (D), day 39 (E), and day 45 (F).
73
Figure 3.7. Chronoamperometry profile and cyclic voltammetry plots of Indium tin oxide
4 electrode, ITO-4. Chronoamperometry profile is shown for the 45-day incubation which
were poised at ~−200 versus SHE (A). Data was recorded at 30 s intervals. Cyclic
voltammetry plots run with a scan range of −600 mV to +300 mV versus SHE with a
scan rate of 1 mV/s: day 7 (B), day 14 (C), day 21 (D), day 39 (E), and day 45 (F).
74
Figure 3.8. Chronoamperometry profile and cyclic voltammetry plots of Graphite 1
electrode (GRA-1). Chronoamperometry profile is shown for the 45-day incubation which
were poised at ~−200 versus SHE (A). Data was recorded at 30 s intervals. Cyclic
voltammetry plots run with a scan range of −600 mV to +300 mV versus SHE with a
scan rate of 1 mV/s: day 7 (B), day 14 (C), day 21 (D), day 39 (E), and day 45 (F).
75
Figure 3.9. Chronoamperometry profile and cyclic voltammetry plots of Graphite 2
electrode (GRA-2). Chronoamperometry profile is shown for the 45-day incubation which
were poised at ~−200 versus SHE (A). Data was recorded at 30 s intervals. Cyclic
voltammetry plots run with a scan range of −600 mV to +300 mV versus SHE with a
scan rate of 1 mV/s: day 7 (B), day 14 (C), day 21 (D), day 39 (E), and day 45 (F).
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Figure 3.10. Chronoamperometry profile and cyclic voltammetry plots of Graphite 3
electrode (GRA-3). Chronoamperometry profile is shown for the 45-day incubation which
were poised at ~−200 versus SHE (A). Data was recorded at 30 s intervals. Cyclic
voltammetry plots run with a scan range of −600 mV to +300 mV versus SHE with a
scan rate of 1 mV/s: day 7 (B), day 14 (C), day 21 (D), day 39 (E), and day 45 (F).
77
Figure 3.11. Chronoamperometry profile and cyclic voltammetry plots of Graphite 4
electrode (GRA-4). Chronoamperometry profile is shown for the 45-day incubation which
were poised at ~−200 versus SHE (A). Data was recorded at 30 s intervals. Cyclic
voltammetry plots run with a scan range of −600 mV to +300 mV versus SHE with a
scan rate of 1 mV/s: day 7 (B), day 14 (C), day 21 (D), day 39 (E), and day 45 (F).
78
3.4.2 Changes in in situ microbial activity over time
(2nd incubation)
In our second incubation, we aimed to provide statistical support for the observations of
both oxidative and reductive measurements at DeMMO 4. CV measurements were
performed more frequently during this run to have a better resolution on the microbial
activities occurring over time. Similar to the preliminary incubation, the terminal CV
measurements demonstrated evidence of catalytic activity (Figure 3.4-3.5). Moreover, we
see a temporal change in both the presence and magnitude of the catalytic activity
throughout the incubation (Figure 3.4-3.11). In both ITO-1 and ITO-2 we see current
becoming more positive over time, that is, at the beginning of the experiment negative
currents were observed (indicating electron uptake) and towards the end of the experiment
current became more positive (indicative of electron deposition) (Figure 3.4A-3.5A). This
is corroborated by the cyclic voltammetry measurements, where we see a distinct catalytic
feature in day 14 and day 21 CV measurements and day 45 CV with a lower max current
(Figure 3.4B-3.4F & Figure 3.5B-3.5F). Additionally, this supports our preliminary
measurements showing anodic catalytic waves, which has also been observed in
metal/electrode reducing microorganisms. (Rowe, Abuyen, et al., 2021) A strong s-type
catalytic features were only observed on ITO-1 and ITO-2 during our second incubation,
and these peaked in activity on a similar time frame (Figure 3.4-3.5). Notably, negative
current was only observed for the other ITO replicates, with stronger activity early in the
incubation (within the first two weeks). The graphite electrodes had a mix of anodic and
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cathodic currents. GRA-1 and GRA-2 predominantly demonstrated negative currents,
coinciding with a more enhanced catalytic signal shown in the CV measurements (Figure
3.8A-3.9A). While GRA-3 and GRA-4 started with positive current and stabilized slightly
below 0 µA (Figure 3.10A-3.11A). GRA-3 was unique in that the current initially generated positive currents and eventually drifted to a more negative current on the 12th day
(Figure 3.10A). Looking at the CV measurements, the general upward trend in the
forward scan supports the potential for mineral reducing microbes present in the
environment (Figure 3.10B-F). However, the predominantly negative currents observed
appears to favor electron uptake mechanisms. Compared with GRA-1 and GRA-2, the
smaller overall currents in the GRA-3 and GRA-4 replicates and generation of anodic
current during CV analysis may support the activity of a mixed community of electrode
reducing and electrode oxidizing reactions in these incubations (Figure 3.10B-F & Figure
3.11B-F).
Differences in spatial and temporal patterns of electrochemical activity were
observed, similar trends were also seen across replicate electrode incubations that were
localized in the same fluid chamber in the reactor—specifically ITO-1 and ITO-2, ITO-3
and ITO-4, GRA-1 and GRA-2 and GRA-3 and GRA-4 (Figure 3.4-3.11). That said, it is
not unlikely that this variation we see in our CV measurements is a result of abiotic and
biotic activities combined. For example, the rise and fall of the max currents in the ITO
measurements is perhaps a range of biological activity and inactivity ultimately
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demonstrating this anodic vs cathodic currents. Interestingly, during similar time frames
replicate sets of ITO 4 (Figure 3.7) and GRA 3 (Figure 3.10) electrodes from this
incubation demonstrated variation in the type of activity (anodic or cathodic) and/or
magnitude despite being in different chambers. Additionally, though these results are
indicative of both anodic and cathodic electrochemical activity in situ, the variability
between electrode types and across replicate electrodes were unexpected. (Xu et al., 2016)
While the driver of these patterns is not defined, it is likely linked to the influence
of the microbial community or the environmental conditions near the electrode surface
favoring a type or level of activity. Moreover, despite attempts to normalize fluid flows at
the beginning of the experiment, rates could have changed throughout the course of the
experiment which points out to the heterogeneity in the DeMMO 4 system. The variability
in fluid flow and/or hydraulic residence times could potentially explain some of the
variability observed ultimately speaking to the strong influence of the environmental
conditions on these results. However, from this work alone it is difficult to assess whether
chemical or biological features are driving the observed variation in patterns we observe
across replicates and over time. Notably, a similar range of peak catalytic activity (based
on the first derivatives of the respective CVs) has been observed across these treatments,
as compared with those seen in the February– May incubation: ranging from −200 to −100
mV versus SHE for the ITO anodic enrichments and −300 to −150 mV versus SHE for
the graphite cathodic enrichment with the majority of observations for both near −200
mV versus SHE. Temporal variance in activity, both in terms of type and level, may
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suggest a low abundance or metabolically versatile community structure containing
representative microbes capable of interacting with solid phase minerals. Though the
driver of electrochemical activity in this system is not known, microbial community
analysis was performed at the end of the experimental incubation to help shed light on
the endemic/active microbial community in these replicate experiments.
3.4.3 Similar microbial community composition observed across
treatments
In a Weighted-Unifrac PCoA (Figure 3.12), the microbial communities clustered tightly
into three groups: primary fracture fluid, first incubation (May samples), and second
incubation (October samples). Similar clusters were also observed for the different distance
methods tested, including Bray–Curtis, and Unweighted Unifrac (data not shown). Note
in Figure 3.12A that the cluster from the fracture fluid includes samples across all time
points, demonstrating temporally stable microbial communities. However, despite a
consistent community structure in the fracture fluids, the different incubations selected
for different microbial communities. Temporal differences in active microbial members,
including succession in the biofilm communities, could explain the differences seen across
the different incubations. A similar trend in consistent community structure across biofilm
communities (during a given experiment) was seen in our previous work for electrodes
ranging in voltage (Jangir et al., 2019), and in different mineral types (sand, calcite, and
a range of manganese, iron, or iron–sulfur minerals) incubated at DeMMO sites 1, 3 & 6.
(Casar et al., 2020)
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Figure 3.12. (A) Principal coordinate plot resulting from weighted-Unifrac PCoA analysis of the
resultant microbial communities based on OTUs. Squares represent samples from the May
sampling trip, triangles represent samples from the October sampling trip, and diamonds indicate
filtered water samples taken from the DeMMO 4 borehole. Legend colors indicate water sampling
date, graphite (GRA), indium tin oxide (ITO), sulfur (Sulf), pyrite and quartz source materials.
OC is used to indicated electrodes that were not poised, or open circuit controls. Axis1 [35.2%]
and Axis 2 [21.1%] indicate the percent variation explained. Principal coordinate plot resulting
from weighted-Unifrac PCoA analysis of the resultant microbial communities based on OTUs. (B)
Principal coordinate plot of the weighted-Unifrac PCoA analysis of the October in situ microbial
community samples. Squares represent graphite samples (GRA), triangles represent ITO samples,
diamonds represent mineral samples (sulfur [Sulf], pyrite, and quartz [Qrtz]) and the plus sign
indicates filtered water sampled from the DeMMO 4 borehole. Axis 1 [30.3%] and Axis 2 [16.8%]
signify the percent variation explained.
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Although all reactor samples were treated and processed the same way, we cannot
completely rule out that some of the differences may be due to differences in efficiency of
nucleic acid recovery from filters versus a biofilm sample. In addition, changes in reactor
design between the Feb-May and Aug-Oct incubations may have slightly altered the
selectivity toward a more anaerobic biofilm community. Many of the same taxa were
observed across the different in situ incubations. The dominant taxa, from the order
Thiotrichales which contain many sulfur-oxidizing microbial lineages, were present in
every sample and made up ~16% of the total sequences (Figure 3.13). They were more
dominant in the May incubation (11.95%-75.54% of total communities) than in the
October incubation (4.93%- 47.84%). The May incubation also showed high enrichment
of the Pseudomonadales (4.88%-53.23%), while the October samples were enriched in the
Chromatiales (10.79%-54.22%)—both putative iron-reducing and/or oxidizing taxa. These
comparisons clearly demonstrate the similarity between the relatively abundant taxa in
the poised potential and open circuit controls (and even some mineral-associated
communities; Figure 3.13), suggesting that the dominant microbial signatures are from
the biofilm-forming community (i.e., the organisms that readily attach to surfaces). The
PCoA plots of the specific electrochemical incubations did not reveal a strong clustering
based on electrode or mineral type (Figure 3.12 B). Notably, the addition of geochemical
data and electrochemical data per sample did not help resolve the community clustering
patterns observed (data not shown). This finding is further supported by our previous
observations from different DeMMO sites (Casar et al., 2020; Jangir et al., 2019), and
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Figure 3.13 Relative 16S rRNA abundance for all community samples collected at the
order level. Orders for the top 100 UTOs plotted for clarity. Remaining OTUs were less
than 0.13% abundance of total sequences.
points to variable or differential metabolic activity of the given community, possibly in
response to changes in water chemistry over time, rather than major shifts in the
community as a whole.
Microbial communities were probed for highly enriched taxa and those that were
only present on specific electrodes (Table 3.2). Because normalized or rarefied abundance
data can mask or inflate changes in specific OTU abundance, we used the DEseq2
extension in the Phyloseq package to analyze the raw sequence data. A focused analysis
of the October samples, which contained more replicates than their May counterpart,
revealed that neither material type nor electrochemical condition (poised vs. open circuit)
correlated well with differential OTU abundances. In fact, only eight OTUs showed
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statistically significant differences in enrichment. (Rowe, Abuyen, et al., 2021) To
deconvolute these differences, we compared poised potential versus open circuit sequences
for each electrode material. While the treatment with graphite electrodes did not yield
any significant enrichments, those with ITO contained 10 taxa significantly enriched on
poised potential electrodes and five taxa more enriched in the open circuit controls (Table
3.2). It should be noted, however, that the OTUs enriched on the poised ITO electrodes
accounted for only 1%–3% of the sequences per community. Some of these OTUs are
related to known electrochemically active microorganisms (e.g., Bacilli and
Deltaproteobacteria), but given their low abundances, it is premature to draw strong
Table 3.2 Significantly enriched OTUs on ITO electrodes compared with ITO open
circuit controls as assessed by DEseq2 (McMurdie & Holmes, 2014)
OTU Kingdom Phylum Class Order
log2FoldChange
p value p adj
denovo34448 Bacteria Proteobacteria Deltaproteobacteria Myxococcales −21.8 4.4E−12 1.0E−09
denovo48911 Bacteria Firmicutes Bacilli Lactobacillales −21.2 1.8E−09 1.8E−07
denovo26753 Bacteria Bacteroidetes Bacteroidia Bacteroidales −21.1 1.1E−10 1.8E−08
denovo38173 Bacteria Planctomycetes Planctomycetia Pirellulales −20.6 2.8E−09 1.8E−07
denovo9913 Bacteria Firmicutes Bacilli Lactobacillales −20.6 3.5E−09 1.8E−07
denovo49676 Bacteria [Thermi] Deinococci Deinococcales −20.6 3.1E−09 1.8E−07
denovo30425 Bacteria Firmicutes Bacilli Bacillales −20.5 3.0E−09 1.8E−07
denovo33825 Bacteria Proteobacteria Alphaproteobacteria Sphingomonadales −20.5 3.3E−09 1.8E−07
denovo57196 Bacteria Actinobacteria Actinobacteria Actinomycetales −20.1 6.6E−09 2.8E−07
denovo48158 Bacteria Bacteroidetes Bacteroidia Bacteroidales −20.1 5.6E−09 2.7E−07
denovo31674 Bacteria Acidobacteria [Chloracidobacteria] RB41 9.7 7.4E−06 2.9E−04
denovo51727 Bacteria Bacteroidetes Flavobacteriia Flavobacteriales 10.3 9.2E−05 3.1E−03
denovo51824 Bacteria Proteobacteria Betaproteobacteria Burkholderiales 10.4 6.6E−05 2.4E−03
denovo2870 Bacteria Tenericutes Mollicutes Mycoplasmatales 10.5 1.92E−04 6.1E−03
denovo39844 Bacteria Actinobacteria Actinobacteria Actinomycetales 25.8 1.1E−17 5.0E−15
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conclusions about their contribution to the observed electrochemical activity; cultivation,
isolation, and further characterization are needed. It is possible, for example, that some
of the dominant microbes in this system promiscuously attached to surfaces and were
electrochemically active but were not enriched during the incubations. The data show
that in this system, the ability to attach to surfaces is a strong driver of microbial
community structure and abundance.
3.5 Conclusion
Studying organisms from limited energy environments such as the subsurface not only
shed light on the extent and diversity of life on our planet but likewise provides insights
into the limits of life, the strategies organisms employ to survive in extreme conditions,
and the potential for life in other planets. In this work, we performed long-term
electrochemical cultivation, combining chronoamperometry and cyclic voltammetry
measurements and showed presence of anodic and cathodic reactions occurring at the site.
Catalytic activity observed from these measurements are consistent with pure cultures
such as Geobacter sulfurreducens and Mariprofundus ferrooxidans, respectively. (Marsili
et al., 2010; Summers et al., 2013) That said, the possible influence of abiotic reactions
occurring together with microbial activity is noteworthy. Additionally, other redox
interactions with microorganisms could generate similar measurements without a direct
link to cellular respiration, i.e. flavins, phenazines, and antibiotics. (Bitew & Amare, 2020;
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Marsili et al., 2008) Based on the ORP of the system and the presence of reduced and
oxidized sulfur we further suggest that the microbial-electrode interactions mimic that of
sulfur in addition to the presence of potential redox couples such as hydrogen with sulfate
or oxygen with sulfide. Previous calculations of thermodynamic feasibility of sulfurreducing and sulfur-oxidizing metabolisms support this claim, (Osburn et al., 2014) as
well as microbial community and metagenomic analyses demonstrating potential for these
metabolisms based on phylogeny and gene presence. (Momper et al., 2017; Osburn et al.,
2014, 2020)
The minerology at SURF supports the presence of iron-bearing minerals, which
depending on the mineral type, could also serve as a source or sink for electrons to support
electrochemically active microbes. Other microbial processes, such as respiratory activities
like iron/manganese mineral respiration, could potentially account for the observed
electrochemical activities. As described previously, given the limited characterization of
these processes, both from a genomic and taxonomic perspective, it is challenging to
identify these processes using 16S abundances and/or metagenomic information alone, this
highlights the importance for continued cultivation and characterization of diverse
electrochemically active microbes.
Lastly, we reiterate that the availability of reactive surfaces and the ability of
microbial taxa to attach to these surfaces appear to be the strongest drivers of microbial
community composition. Attachment may offer a strong selective advantage to microbes
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in this environment as under these conditions attachment is likely a ‘first come first serve’
phenomenon. That said, since attachment seems to be the stronger selection factor as to
who ends up on a redox-active surface, it is difficult to conclude on which organisms are
electrochemically active in the community. However, utilization of in situ electrodes for
subsequent laboratory enrichment and isolation can be used to confirm the presence of
electrochemically active organisms in our in situ reactors.
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Chapter 4: Electrochemical
investigation of Aggregatibacter
actinomycetecomitans D7S-1
4.1 Abstract
Aggregatibacter actinomycetecomitans D7S-1 is a pathogen and oral commensal that is
strongly associated with aggressive forms of periodontitis. It has been suggested that it
can perform extracellular electron transfer, but the organism’s electroactivity is poorly
understood. In this study, we used amperometric and voltammetric measurements to
investigate in vivo electrochemical activity. Here, we illustrated a reduction in anodic
current upon the introduction of fresh media, followed by a recovery of current upon
reintroducing cell-free spent media. Notably, voltammetry analysis on cell-free spent
media indicated redox-active material with a redox potential at around -300 mV (vs SHE).
Tests on candidate electron mediator, FMN, did not increase anodic current in cellcontaining reactors relative to sterile control reactors. Moreover, chemical inhibition of ctype cytochromes, using KCN and NaN3 showed no effect on current generated from cell
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addition. Notably, anodic current comparison between live and dead (heat-killed) cells
demonstrated statistically similar current levels. In summary, this study showed that in
the conditions tested, Aggregatibacter actinomycetecomitans D7S-1 is not
electrochemically active and cannot perform extracellular electron transfer.
4.2 Introduction
Extracellular electron transfer (EET) is a metabolic process by which bacteria use the
electrons derived from the oxidation of an electron source such as lactate or acetate and
transfer the electrons to minerals such as iron oxides, manganese oxides or electrodes
outside of the cell. (Myers & Nealson, 1988; Pirbadian et al., 2020) While the diversity of
EET capable bacteria are growing, Geobacter sulfurreducens and Shewanella oneidensis
are the most characterized metal oxide reducers. (C. H. Chan et al., 2017; Chong et al.,
2022; Pirbadian & El-Naggar, 2012; Zacharoff et al., 2017) The number of bacteria from
the human microbiome that are known to be capable of EET is a relatively short list
compared to those that are found in the environment. (Jangir et al., 2016; Karbelkar et
al., 2019; Pérez-Rodríguez et al., 2021; Rowe et al., 2019) Given the complexity of the
human microbiome, with the diverse microbial community interacting with one another
and with the host, developing strategies to study this microbial ecology and the factors
influencing the microbial community is an important challenge that needs to be
investigated. One way of studying these organisms is via electrochemical methods. In this
instance, instead of using iron/manganese oxides as the electron acceptor, a poised
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electrode is used, mimicking the redox potential of the oxides. This method allows for
real-time monitoring of electrochemical parameters such as current and potential.
Additionally, this allows researchers to monitor the production or consumption of specific
metabolites.
Human-health related bacteria that have been investigated for their EET capability
include, Listeria monocytogenes and Enterococcus faecalis. L. monocytogenes is a Grampositive bacterium discovered to perform EET through the release of electron shuttles
outside of the cell. (Light et al., 2018) L. monocytogenes was first discovered to reduce
ferric iron via a simple ferric reductase plate assay in 1993 and it wasn’t until 2018 that
it was shown to be electrochemically active transferring electrons via a flavin-based EET
mechanism. (Deneer & Boychuk, 1993; Light et al., 2018) Studies on another Grampositive bacterium, E. faecalis, demonstrated the relevant role of the quinone pool in the
ability of E. faecalis to perform EET. (Pankratova et al., 2018) Not surprisingly, both
Gram-positive E. faecalis and L. monocytogenes share homologous structures that
facilitate the flavin-based EET mechanism. With these two examples, it is clear that
electrochemical investigations on new organisms pave the way to the discovery of new
mechanisms, expanding scientific knowledge and filling in knowledge gaps ultimately
moving the field forward. This is especially true for the human microbiome and pathogens
as it is less understood compared to environmental microbes. Additionally, pathogenic
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organisms have a significant impact on human health and is essential for preventing and
controlling infectious diseases.
Organisms from the human oral microbiome have also been isolated and proposed
to perform EET. Oral pathogenic strains and members of polymicrobial biofilms such as
Corynebacterium matruchotii, Streptococcus mutans, Porphyromonas gingivalis, and
Aggregatibacter actinomycetecomitans have been shown to produce modest current in
electrochemical cells. (Miran et al., 2021; Naradasu et al., 2020) These organisms however
produce relatively low currents compared to the well characterized counterparts including
S. oneidensis, and even compared to L. monocytogenes and E. faecalis. (Light et al., 2018;
Pankratova et al., 2018; Pirbadian et al., 2020) These low current levels in turn bring up
concerns as to whether these organisms are truly EET active. Notably, the definition of
electrochemical activity is not fully defined, there is no threshold as to what constitute as
biological extracellular electron transfer.
In this work, we investigate the capability of Aggregatibacter
actinomycetecomitans D7S-1 to perform EET. A. actinomycetecomitans strain Y4 has
been shown to be electrochemically active, producing current in anodic conditions.
Naradasu and coworkers indicated that according to their redox-dependent 3,3'-
diaminobenzidine (DAB) chemical staining of cell membranes, there is significant cellsurface bound redox protein. They also proposed EET shuttling mechanism involved via
their cell-free voltammetry analyses. (Naradasu et al., 2020) Voltammetry itself, however,
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is not a definitive measurement to determine shuttling, it needs to be coupled with other
measurements, such as media exchange where one removes released molecules and observe
decrease in EET activity. (Karbelkar et al., 2019; Lam et al., 2019)
While Naradasu et al. provided a good foundation, further investigations are
necessary in order to fully understand the mechanism underlying the electrochemical
activity observed in A. actinomycetecomitans. The present study employed
electrochemical measurements to potentially shed some light on the EET mechanisms
involved in A. actinomycetecomitans D7S-1. Here, by performing media exchange
experiments followed by reintroduction of the spent media, we show that A.
actinomycetecomitans D7S-1 produces soluble redox active molecules that interacts with
the anode, generating low current levels. This led us to investigate the possible role of
FMN as an electron shuttle and found no definitive influence on A. actinomycetecomitans
D7S-1 ability to perform EET. Additionally, we looked into the effects of cytochrome
inhibitors and likewise performed heat-killed cell electrochemical assays and found no
difference between pre- and post-inhibition. While we observe change in current during
media exchange, findings from our other in vivo assays, such as the inhibition assays,
point to the electrochemical inactivity of A. actinomycetecomitans D7S-1.
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4.3 Methods
4.3.1 Cell Growth Conditions
A. actinomycetecomitans D7S-1 was grown in Brucella blood agar plates (Hardy
Diagnostics) incubated in microaerobic conditions under 95:5 (v/v) N2/CO2 atmosphere
at 37 °C for 48 h. Colonies were picked to inoculate a 25 mL microaerobic (5 mL filtered
air) 95:5 (v/v) N2/CO2 TSBYE media: 17 g/L Tryptone, 2.5 g/L K2HPO4, 2.5 Glucose,
5 g/L NaCl, 3 g/L Soytone, 6 g/L Yeast Extract, pH 7.3 ± 0.2. Liquid cultures were
incubated at 37° C for 48 h shaking at 160 RPM. For reactor inoculation, cells were first
washed with a defined media (DM) 3X to remove redox active compounds. This was
followed by resuspension in 1 mL DM-anode. DM contained (per liter): 0.2 g MgCl2*6H2O,
1.0 g NH4Cl, 0.2 g CaCl2*2H2O, 0.9 g NaCl, 2.5 g NaHCO3, 7.2 g HEPES buffer. The pH
of the medium was adjusted to 7.0 using NaOH. Reactor medium (DM-anode) is composed
of DM with the addition of 30 mM DL-lactate and amino acids for anodic cultivation at
+435 mV vs SHE. Working electrode potential was selected based on the potential used
in a prior study conducted on A. actinomycetecomitans. (Naradasu et al., 2020)
4.3.2 Reactor setup and electrochemical measurements
A custom-built single chamber bioreactor was used to investigate the electroactivity of A.
actinomycetecomitans. For chronoamperometry measurements (+435 mV vs SHE), the
bioreactor was composed of a graphite felt electrode (POCO AXF-5Q 0.059” x 0.225” x
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0.83”) (Tri-Gemini LLC, CAT-num: XM15839C), coiled platinum wire served as the
counter electrode, and an Ag/AgCl in 1M KCl reference electrode (CH Instruments, Inc.).
To prepare the working electrode it was kept in 1 M HCl for 5 hours then rinsed with deionized water 10 times and sonicated twice, first in distilled de-ionized water, then in
ethanol. The working electrodes were stored in DI water until further processing. After
reactor assembly, the reactor is kept in 1 M HCl to further ensure the absence of trace
metals that could potentially cause background signal. This is done for about 5 hours and
then rinsed 10X with Milli-Q H2O to full volume and then autoclaved. All electrochemical
analysis were performed under nitrogen atmosphere, at 37 °C using a CHI1000 8-channel
potentiostat (CH Instruments, Inc.). Baseline current level was set for a few hours before
reactors were inoculated with cells to a final OD of 0.05.
Flavin mononucleotide (FMN) has been shown to improve anodic current in other
electroactive bacteria. (Okamoto, Hashimoto, et al., 2014; von Canstein et al., 2008; Xu
et al., 2016) To that end, we investigated the influence of FMN in the electroactivity of
A. actinomycetecomitans. To test this, we started the experiment by first allowing the
biofilm to colonize the electrode by poising the electrode at +435 mV vs SHE for about
20 hours. 200 nM FMN was added followed by another 400 nM FMN (final concentration
of 600 nM) during a continuous monitoring of current generation.
To investigate whether the current generated is biological, we performed inhibitory
electrochemical assays. For the chemical inhibition assay here we again let the cells form
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a biofilm by poising the electrode at +435 mV vs SHE for about 20 hours. After which
we added 5 mM potassium cyanide (KCN), then after about 4 hours we added 10 mM
sodium azide (NaN3) and continued measuring current. Physical inhibition was also
performed. First, we grew A. actinomycetecomitans D7S-1 in TSBYE as mentioned
earlier. After 48 h of incubation cultures were incubated at 50 °C for 2 h shaking at 160
RPM. Cultures were then washed prior to reactor inoculation and chronoamperometry
measurements.
4.3.3 Media exchange and spent medium analysis
Electrode-attached biofilms were further analyzed to determine whether EET is happening
via direct cell-to-electrode contact or through soluble electron shuttles. In brief, after 20
h of anodic cultivation at +435 mV (vs SHE) reactors were taken to the anerobic chamber
where the WE colonized by A. actinomycetecomitans D7S-1 was transferred to a fresh
reactor containing media with 30 mM lactate then chronoamperometry measurements
were continued. The supernatant from the first incubation was processed to separate the
planktonic cells from the medium (2500 RPM for 30 minutes, repeated 3 times) after
which the spent media was made anaerobic by bubbling N2 for 30 minutes. After a few
hours of electrocultivation in the fresh medium the WE electrode was re-introduced to the
spent media, and chronoamperometry measurement was continued.
To perform electrochemical measurements on the spent media another set of the
resulting anaerobic spent medium was transferred to a new electrochemical reactor for
97
scan rate dependence cyclic voltammetry measurements. To improve the sensitivity of the
measurements, a 3 mm diameter Gold (Au) Electrode (BASi Inc.) was used as the working
electrode. Platinum wire and Ag/AgCl in 1 M KCl was used as counter electrode and
reference electrode, respectively. Measurements were done in anaerobic conditions
scanning a potential range of -700 mV to +200 mV vs SHE at scan rates of 100 mV/s, 50
mV/s, 10 mV/s & 1mV/s.
4.3.4 SEM Imaging
After 20 h of anodic cultivation graphite felt working electrodes were processed for
Scanning Electron Microscopy (SEM). First, the electrodes were fixed using 2.5%
glutaraldehyde for about 20 h. Samples were then subjected to an ethanol dehydration
process (30, 50, 70, 80, 90, 95 and 100% v/v ethanol) 3x for 10 min of each ethanol
concentration before moving to the next higher concentration. The samples were kept in
the desiccator for 24 h until further processing. For SEM imaging, samples were mounted
on aluminum stubs and sputter coated with carbon (Sputter coater 108, Cressington
Scientific), followed by imaging at 3 keV using a JEOL JSM 7001F low vacuum field
emission SEM.
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4.4 Results
In this work we set out to further electrochemically characterize A. actinomycetecomitans
D7S-1. (Tjokro et al., 2019) It has been shown that lactate is an important carbon source
for A. actinomycetecomitans, (Brown & Whiteley, 2007, 2009) moreover, previous
electrochemical investigations point to the importance of lactate in the extracellular
electron transfer activity of the organism as well. (Naradasu et al., 2020) To that end, we
looked into the sequence similarity of FMN-dependent lactate dehydrogenase (lldD) in
strains D7S-1 and Y4 using NCBI blast. Our analysis showed a 99.7% sequence similarity
with a 100% query coverage.
Figure 4.1. A. actinomycetecomitans produces soluble electron shuttles.
Chronoamperometry measurement on A. actinomycetecomitans with the working
electrode poised at +435 mV (vs SHE) with 30 mM DL-lactate. Arrow heads indicate
inoculation time, fresh media exchange and reintroduction of spent media. Reactors were
kept in nitrogen atmosphere at 37 °C
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4.4.1 Replacement of reactor media abolishes Aggregatibacter
actinomycetecomitans anodic current
To characterize the electrochemical activity of A. actinomycetecomitans D7S-1 we used a
three-electrode electrochemical reactor containing 30 mM DL-lactate as the electron
source and the working electrode poised at +435 mV (vs SHE) as the electron acceptor.
Upon the addition of the cells, chronoamperometry measurements showed a rise in current
level, suggesting possible electrochemical activity of the organism (Figure 4.1). This is
consistent with previous work. (Naradasu et al., 2020) Next, we set out to investigate the
possible involvement of soluble electron shuttles. It has been suggested that A.
actinomycetecomitans might have the ability to produce redox-active soluble electron
shuttles. (Naradasu et al., 2020) It is likely then that A. actinomycetecomitans D7S-1
Figure 4.2. A. actinomycetecomitans robust biofilm on the electrode. Scanning Electron
Microscopy (SEM) of A. actinomycetecomitans biofilm on graphite felt electrode fibers.
(A) Image at 10,000X magnification (10 µm scale bar). White rectangle indicate image in
B. (B) Image at 65,000X magnification (1 µm scale bar).
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could be generating or releasing some redox-active molecules that can produce such
current. To investigate this, we performed a media exchange experiment. Here we let the
biofilm colonize the electrode for about 20 h on an anode which was then followed by
replacing the reactor (‘spent’) medium with fresh medium. Media replacement eradicated
more than 50% of the current generated (Figure 4.1). Subsequent reintroduction of the
cell-free spent media recovered most of the current back. This recovery of current upon
reintroduction of spent media is consistent with investigations on other bacterial species
that produce soluble redox shuttles for electrochemical activity. (Karbelkar et al., 2019)
Given we used a graphite felt electrode, which is very absorbent, it is likely that some of
the redox-active molecules were not removed from the system which would then explain
Figure 4.3. Scan rate dependence of the cyclic voltametric response of spent media A.
actinomycetecomitans. (A) Cyclic voltammetry measurement of reactor spent media. The
potential was swept -700 mV to 200 mV vs SHE at 1 mV/s, 10 mV/s, 50 mV/s and 100
mV/s. inset shows voltammetry measurements of sterile media. (B) Peak current vs scan
rate plot.
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the limited eradication of current upon the introduction of the fresh media. However, the
data from this media exchange experiment does still suggest presence of soluble redoxactive molecules in solution. Following chronoamperometry measurement we imaged the
biofilm attached to the electrode through a Scanning Electron Microscope (SEM) and saw
robust biofilm formations (Figure 4.2).
4.4.2 Spent media contains redox-active molecules
Further analysis of the spent medium using a more sensitive electrode was conducted to
shed light on the electrochemical property of the molecule present in solution. Cyclic
voltammetry measurements on the isolated spent medium shows a peak around -300 mV
(vs SHE) (Figure 4.3) We further characterized the cell-free spent media scanning a range
of potential at different scan rates; 1 mV/s, 10 mV/s, 50 mV/s, and 100 mV/s. Peak
current observed around -300 mV increased with increasing scan rates. This analysis
suggests the presence of a soluble molecule that’s capable of interacting with the electrode.
4.4.3 Flavin mononucleotide has no effect on the EET activity of
A. actinomycetecomitans
The measured redox potential from the voltammetry analysis is relatively close to the
redox potential of flavin mononucleotide (FMN). (Xu et al., 2016) Consequently, we
proceeded to perform chronoamperometry measurements with the addition of FMN into
cell-containing reactors. Upon the addition of 200 nM FMN no significant influence was
measured on the cell containing reactors while an obvious increase in current was observed
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Figure 4.4. Addition of FMN does not improve anodic current relative to the negative
sterile control. Chronoamperometry measurement on A. actinomycetecomitans with the
working electrode poised at +435 mV (vs SHE). Arrow heads indicate addition of 200 nM
and 400 nM FMN. Reactors were kept in nitrogen atmosphere at 37 °C.
in negative (i.e. cell-free, abiotic) controls (Figure 4.4). Addition of 400 nM FMN resulted
in an even greater current change in the negative controls while the cell containing reactors
had a moderate increase. The higher current change observed in the negative control can
perhaps be explained by the surface area available for FMN to interact with; the abiotic
electrodes have a higher electrochemically active surface area for interaction with the
medium while cells cover a significant fraction of the biotic electrodes.
4.4.4 Chemical and physical inhibition has no effect on anodic
current production of A. actinomycetecomitans
To investigate whether the anodic current observed in A. actinomycetecomitans D7S-1
reactors is of biological origin, we performed chemical inhibition assays to target some
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cytochromes that can potentially be involved in the EET pathway in A.
actinomycetecomitans D7S-1. Potassium cyanide (KCN) and sodium azide (NaN3) inhibits
cellular electron transfer by binding to heme groups, ultimately preventing redox reactions
from occurring and thus halting important constituents of electron transfer chains.
(Karbelkar et al., 2019; Léger & Bertrand, 2008; Rowe et al., 2018; Woźnica et al., 2003)
During a chronoamperometry measurement, addition of 5 mM KCN at first brought the
current to and eventually stabilized at above cell baseline levels (Figure 4.5). This rise
can be attributed to the electrochemical oxidation of KCN to cyanate (OCN-), a less toxic
product. (Léger & Bertrand, 2008) However, the concentration added to the reactor is
relatively high ensuring cells will be exposed to the inhibitor. Nevertheless, we followed
the KCN inhibition with the addition of 10 mM NaN3 (Figure 4.5). Here we see that NaN3
Figure 4.5. A. actinomycetecomitans is not sensitive to cytochrome inhibitors KCN and
NaN3. Chronoamperometry measurement on A. actinomycetecomitans with the working
electrode poised at +435 mV (vs SHE). Arrow heads indicate addition of 5 mM KCN and
10 mM NaN3. Reactors were kept in nitrogen atmosphere at 37 °C.
104
addition had no effect on the anodic current/did not eradicate the current below the
baseline prior to addition of inhibitors. Given that KCN and NaN3 inhibit specific electron
transport chain proteins, it is possible that we simply did not use the appropriate
inhibitors. To that end, we performed an experiment where we compared the
electrochemical activity of live and heat-killed cells (2 h incubation at 50 °C). Without
heat treatment cell viability was around 5.7×1011 CFU/mL while the heat-killed culture
retained 5×103 CFU/mL live cells. Surprisingly, the anodic currents observed from these
two conditions are statistically similar, live cells had current levels at 0.543 ± 0.17 µA
while heat-killed cells had 0.438 ± 0.11 µA (Figure 4.6). To investigate the influence of
the cell suspension solution to the current change upon inoculation, we inoculated the
Figure 4.6. Current levels between Live and Heat-killed A. actinomycetecomitans are
comparable. Average anodic current levels between Live and Heat-killed cells. Error bars
represent standard error of triplicate experiments. Working electrode poised at +435 mV
(vs SHE) with 30 mM DL-lactate. Reactors were kept in nitrogen atmosphere at 37 °C.
105
reactors with just the supernatant from the cell suspension after washing the cells three
times. (Data not shown) Here we saw no difference between before or after addition of
the supernatant, current remained at baseline levels.
4.4.4 An OMC Shewanella mutant has higher anodic current
compared to A. actinomycetecomitans D7S-1
We also compared the anodic current production of A. actinomycetecomitans to an
electrochemically inactive S. oneidensis mutant (∆OMC) lacking the omcA, mtrA, mtrF,
dmsE, SO4360, cctA, and recA genes, as well as the known electrochemically active wildtype S. oneidensis MR-1. While WT S. oneidensis produced approximately 20 µA of
current in 12 hours, A. actinomycetecomitans D7S-1 showed a current level even lower
than ∆OMC, nearly reaching 0 µA, which is close to the current levels observed in the
blank reactor (Figure 4.7). Looking into the genome, we used FeGenie, a software program
that allows scientific investigators to perform a preliminary check on the presence of ironrelated genes such as canonical iron reductase/oxidase (from metal reducers/oxidizers),
iron import/export proteins, siderophore import/export and synthesis, and iron storage.
(Garber et al., 2020) Using this analysis, we found no indication of iron reductase genes
present in either D7S-1 and Y4 strains which is consistent with our electrochemical
measurements. Taken together, the current observed in A. actinomycetecomitans D7S-1
are insignificant levels and perhaps the organism should be categorized as
electrochemically inactive, contrary to recent reports.
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Figure 4.7. A. actinomycetecomitans generates anodic current lower than S. oneidensis
∆OMC mutant. Chronoamperometry experiments (+435 mV vs SHE) of S. oneidensis
WT, outer membrane cytochrome (OMC) mutant (ΔomcA/ΔmtrA/ΔmtrF/ΔdmsE/
ΔSO4360/ΔcctA/ΔrecA), A. actinomycetecomitans (Aa) and sterile control reactor with
20 mM sodium lactate.
4.5 Discussion
The Gram-negative commensal bacterium Aggregatibacter actinomycetecomitans has
been suggested to be capable of extracellular electron transfer (EET). (Naradasu et al.,
2020) In our work, we found contradicting data, suggesting that the current observed in
A. actinomycetecomitans is not related to EET. This we show through our inhibition
assay via the addition of c-type cytochrome inhibitors KCN and NaN3, where we observed
no indication of current decrease in the cell-containing reactors upon the addition of the
inhibitors. Given we only tried two inhibitors, we likewise performed experiments on heatkilled cells, here we observed live and heat-killed cells with similar current levels. Taken
together, the absence of any impact on the current production due to addition of
107
inhibitors, and the similar currents production observed in heat-killed control experiments,
suggests that the anodic current we observed in our early measurements is unlikely to be
of biological origin. In this chapter we also show that addition of FMN has no effect on
the anodic current relative to the sterile negative controls. With that said, the genome of
A. actinomycetecomitans shows that it is capable of producing flavins. (data not shown)
Given we did not explore alternative electron shuttles, further investigation into the role
of electron shuttles is warranted. Moreover, electrode material can also influence signal
acquisition in the presence of flavins. Glassy ITO electrodes tend to have a low affinity
for flavin adsorption, high surface area carbon cloth or graphite felt on the other hand not
only allows for biofilm formation, but likewise have a greater affinity for flavins which in
this study was illustrated by the increased current response from sterile reactors. (Xu et
al., 2016)
One possible mechanism by which A. actinomycetecomitans may function is by
supplying flavins for utilization by other microbes. Microbial consortia typically consist of
organisms that collaborate, with one species supplying specific nutrients or molecules for
the benefit of others. Production of lactate by lactic acid bacteria provide carbon source
for multiple organisms including A. actinomycetecomitans. (Ramsey et al., 2011) S.
oneidensis and E. coli is another example, where flavins generated by S. oneidensis, is used
for extracellular electron transfer by E. coli. (Wang et al., 2015) In our study, we showed
that A. actinomycetecomitans D7S-1 produces a redox-active material. (Figure 4.1) That
108
said, it is possible that A. actinomycetecomitans D7S-1 is not capable of performing EET,
but can generate electron shuttles for its neighbors.
While the field has focused on metal reducers and oxidizers as models for EET,
there has been a rapid increase of studies reporting electrochemical activity in a wide
range of organisms. Some of these recent reports describe organisms with very low
electrochemical activity and where it is unclear why/how they perform EET. This we
refer to as weak electricigens, microorganisms that exhibit relatively low levels of
electrochemical activity compared to organisms like S. oneidensis. It is important to note,
however, that these low electrochemical activities do not necessarily mean the organism
is able to perform EET, that is the electrochemical activity is not linked to respiration or
metabolism in general. A. actinomycetecomitans would fit this definition based on
previous reports. We replicated these reports as shown in Figures 4.1, 4.3 & 4.5 however,
we subsequently discovered that the extremely low currents observed persisted even when
inhibitors were present and when cells were subjected to heat prior to the measurement.
4.6 Conclusion
Extracellular electron transfer (EET) between microorganisms and solid electron
acceptors/donors, such as electrodes and iron minerals, is a prevalent phenomenon with
significant implications for biogeochemical cycles, microbial ecology and now in the human
microbiome. Consequently, there is a widespread interest in elucidating the mechanisms
109
by which microorganisms conduct electron transfer between intracellular and extracellular
environments. For example, EET has been hypothesized to play a critical role in
supporting growth in the intestinal lumen considering the significance of anaerobic growth
mechanisms for microbial proliferation. (Light et al., 2018) Recently, a number of
unconventional electroactive microorganisms have been proposed to be electrochemically
active. These microorganisms are currently being classified as weak electrogenic bacteria
or weak electricigens, current signals from which are low and unpredictable. (Aiyer &
Doyle, 2022; Doyle & Marsili, 2018) According to the results described here, current
generated by Aggregatibacter actinomycetecomitans D7S-1 is low, unpredictable and
perhaps a result of abiotic reactions. We showed that there are endogenous redox active
soluble materials released by A. actinomycetecomitans, however the absence of current
amplification upon the addition of exogenous flavins suggests that the organism does not
use the electron shuttles for EET purposes (Figure 4.4). Additionally, to determine
whether the current is biological we performed inhibition assays via chemical and physical
approaches. Nevertheless, current remained above or equal to preinhibition conditions
(Figure 4.5 & 4.6).
Results from this work also shows that A. actinomycetecomitans has current levels
below S. oneidensis ∆OMC. In the field of Microbial Electrochemistry, particularly for
those who work with S. oneidensis MR-1, ∆OMC is used as the electrochemically inactive
negative control, prompting the question whether anodic currents below these thresholds
110
should be regarded as abiotic. (Abuyen & El-Naggar, 2023; Ross et al., 2011) As research
in this emerging field progresses, the long-term consequences and potential implications
of these unexpectedly electroactive microorganisms will become more evident. It is then
necessary to define biological electrochemical activity, perhaps by relating it to the
magnitude of energy flux (power) to keep an organism active.
111
Chapter 5: Conclusions
Microbial diversity plays a significant role in driving extracellular electron transfer (EET)
processes in various ecosystems, such as soils, aquatic environments, deep subsurface
terrestrial environments, and the human microbiome. (Bar-On et al., 2018; Casar et al.,
2020; LaRowe & Amend, 2015; Magnabosco et al., 2018; Osburn et al., 2014) Moreover,
microbes have the capacity to employ diverse electron acceptors for anaerobic respiration,
depending on their metabolic capabilities and the prevailing environmental factors. This
versatility enables them to adjust to fluctuations in environmental conditions.
In Chapter 2, an electrochemical investigation on the role of soluble FeCl2 in the
extracellular electron uptake (EEU) by Shewanella oneidensis MR-1 was discussed. Here
it was demonstrated that FeCl2 can increase EEU in S. oneidensis in an electrochemical
reactor with fumarate serving as the intracellular electron acceptor. The FeCl2
concentration-dependent electron uptake from cathodes ceased when an iron chelator was
introduced. Moreover, S. oneidensis was only able to take up electrons from the cathode
in the presence of fumarate reductase (fccA), confirming that electrons from the cathodes
enters the periplasm. Additionally, mutants of S. oneidensis lacking certain outer
112
membrane and periplasmic cytochromes showed notably reduced current levels compared
to the wild type strain. These findings suggest that soluble iron can serve as an electron
transporter to the extracellular electron machinery of S. oneidensis.
In Chapter 3, an extended electrochemical cultivation conducted at Sanford
Underground Research Facility (SURF) in South Dakota is discussed. Here
chronoamperometry and cyclic voltammetry techniques were used, revealing both anodic
and cathodic reactions occurring at the site. Catalytic reactions observed from cyclic
voltammetry measurements in both anodic and cathodic conditions align with
observations from pure cultures such as Geobacter sulfurreducens and Mariprofundus
ferrooxidans, respectively. While potential influence of abiotic reactions can occur
concurrently with microbial activity in this environment, other redox interactions
involving flavins and phenazines could yield similar measurements. However, considering
the oxidation-reduction potential (ORP) of the system coupled with the presence of sulfur
species at the site, we suggest that the microbial interactions we are seeing in our
measurements mirror microbe-mineral interactions. This proposition is supported by
previous thermodynamic feasibility calculations, as well as microbial community and
metagenomic analyses. (Momper et al., 2017; Osburn et al., 2014) However, it is
emphasized that the availability of reactive surfaces and capacity of microbial taxa to
attach to these surfaces appear to be the primary determinant of microbial community
composition. That said, it is challenging to conclusively identify which organisms are
113
electrochemically active within the community. Future research in this area could focus
on isolation and electrochemical characterization of organisms from this site to validate
the findings discussed in this chapter.
In Chapter 4, we find that the current observed upon in Aggregatibacter
actinomycetecomitans D7S-1 is not due to extracellular electron transfer does not exhibit
extracellular electron transfer capabilities. Instead, we observed the release of endogenous
redox-active soluble materials by A. actinomycetecomitans. The redox potential of these
molecules closely resembles that of FMN, prompting an investigation into the potential
involvement of FMN in the anodic current generation in A. actinomycetecomitans.
However, our experimental results did not provide conclusive evidence of flavin-mediated
extracellular electron transfer. Furthermore, inhibition assays using chemical and physical
methods were conducted to assess the biological origin of the observed current. Despite
these efforts, the current persisted at level equal to than those observed before inhibition,
suggesting that A. actinomycetecomitans is unlikely to be electrochemically active.
In conclusion, the studies discussed in this thesis underscore the critical role of
microbial diversity in driving extracellular electron transfer processes across various
ecosystems. The versatility of the microorganisms in utilizing diverse electron acceptors
for anaerobic respiration further emphasizes their adaptability to changing environmental
conditions. The findings presented here illustrates specific instances of microbe-electrode
114
interactions and anerobic respiratory pathways, shedding light on the complexities of
microbial metabolism and environmental redox processes.
115
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_0005.JPEG
Abstract (if available)
Abstract
Extracellular electron transfer (EET) is a respiratory strategy that allows bacteria to access external electron acceptors such as redox-active elements (e. g., Fe, Mn) in solid minerals. Extracellular electron uptake (EEU), on the other hand, refers to the capacity of microorganisms to acquire electrons from solid-phase conductive materials like metal oxides, a less understood mechanism. Organisms such as Shewanella oneidensis and Geobacter sulfurreducens are two of the well-studied organisms capable of EET. In S. oneidensis, the oxidation of a carbon source occurs in the cytoplasmic space and gained electrons are transferred from heme containing inner membrane proteins, through the periplasm and eventually to the outer membrane reducing external electron acceptors. To date there are three known ways S. oneidensis respire external electron acceptors, by direct contact of outer membrane cytochromes, through soluble electron shuttles, and lastly via membrane extensions containing outer membrane cytochromes. Much of our mechanistic understanding of EET is derived from studies of transmembrane cytochrome complexes and extracellular redox shuttles that mediate outward EET to anodes and external electron acceptors. In contrast, there are knowledge gaps concerning the reverse process of inward EET (EEU) from external electron donors to cells. To address this knowledge gap, the first project discussed in this dissertation touches on the role of soluble iron in enhancing EET from cathodes to the model EET bacterium S. oneidensis MR-1. Here we use amperometric and voltammetric tools coupled with gene deletions to demonstrate that S. oneidensis can use FeCl2 as an electron shuttle to enhance cathodic electron uptake. Our data reveals that FeCl2 does enhance electron uptake by S. oneidensis with the involvement of the Mtr pathway all the way to the periplasm reducing electron acceptor fumarate to succinate.
The second project we discuss in this dissertation looks into how EET may contribute in maintaining microbial communities in the deep-subsurface terrestrial environments. At the Deep Mine Microbial Observatory (DeMMO) located in Sanford Underground Research Facility (SURF) in Lead, South Dakota, we conducted long-term in situ electrochemical experiments aimed at evaluating the capability of deep groundwater microbial communities to employ extracellular electron transfer to support microbial respiration. We couple electrochemical measurements with 16S rRNA analysis to look at the influence of poised electrodes on the microbial population on a conductive surface. Our results indicate that there are anodic and cathodic activities at the site. Moreover, the ability to attach to surfaces is the main driver of microbial composition on minerals and electrodes.
In the last chapter of this dissertation, we focus on the human microbiome where the number of bacteria that are known to be capable of EET is relatively small compared to those that are found in the environment. Specifically, we performed electrochemical investigations on a microbial isolate from the oral microbiome, Aggregatibacter actinomycetecomitans, previously proposed to be capable of EET. Aggregatibacter actinomycetecomitans D7S-1 represents a significant threat to human health, being a Gram-negative opportunistic pathogen and a member of the oral microbiota strongly associated with severe types of periodontal disease. Amperometric and inhibition assays were conducted to investigate the extent of microbial electrochemical activity of this strain. While the inoculation of electrochemical reactors with A. actinomycetecomitans results in weak current production, our study highlights the importance of controls designed to test whether the observed current in such ‘weak electrogens’ stems from in vivo activity.
In sum, we discovered an inward Fe-mediated extracellular electron transfer (EET) mechanism in a well-studied EET model organism marked a significant breakthrough. Additionally, we expanded electrochemical studies to include “non-canonical” EET microbes, both in their natural environment and within the human microbiome. This expansion broadened our understanding of microbial electrochemical interactions and their potential roles in various ecosystems and human health.
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Abuyen, Karla
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Core Title
Electrochemical studies of outward and inward extracellular electron transfer by microorganisms from diverse environments
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College of Letters, Arts and Sciences
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Doctor of Philosophy
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Molecular Biology
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2024-08
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05/28/2024
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anaerobic respiration,electrode oxidation,electrode reduction,electron shuttle,environmental microbiology,extracellular electron transfer,extracellular electron uptake,human microbiome,metal oxidation,metal reduction,microbial electrochemistry,OAI-PMH Harvest
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Tags
anaerobic respiration
electrode oxidation
electrode reduction
electron shuttle
environmental microbiology
extracellular electron transfer
extracellular electron uptake
human microbiome
metal oxidation
metal reduction
microbial electrochemistry