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Electrochemical investigations and imaging tools for understanding extracellular electron transfer in phylogenetically diverse bacteria

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Electrochemical investigations and imaging tools for understanding extracellular electron transfer in phylogenetically diverse bacteria

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Content ELECTROCHEMICAL INVESTIGATIONS AND IMAGING TOOLS FOR
UNDERSTANDING EXTRACELLULAR ELECTRON TRANSFER IN
PHYLOGENETICALLY DIVERSE BACTERIA
by
Amruta Karbelkar
A Dissertation Presented to the
FACULTY OF THE GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulllment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(CHEMISTRY)
December 2019
Copyright 2019 Amruta Karbelkar
to those who keep going
ii
Acknowledgments
First of all, I would like to thank my advisor, Moh El-Naggar. I feel so privileged
to have been a part of your lab. You have been an excellent advisor, a great teacher
and a patient mentor. I have not only grown as a scientist, but also as a human
being under your guidance. You have encouraged me to go out of my comfort zone
and provided me with all the support and resources necessary to make it happen.
As a result, I have been able to explore an exciting subject and meet amazing
people. I am forever indebted to you, for everything you have done for me.
I am so thankful to have had the opportunity to collaborate with Dr. Steve
Finkel. Steve, you always made me feel like I was a part of your group. After
Moh's lab, your lab was like a second home to me. Whether it was you guiding
me on a project or us having a chat in the breakroom, I will always cherish all the
time I spent with you. Thank you so much, Steve.
I want to thank the current and past El-Naggar lab members { the most won-
derful blend of people I have ever worked with! Sahand, Matt, Lori, Annie, Hyesuk,
Grace, Marko, Karla, Christina, Nicole and Leila { you all made sure that the lab
was always full of enthusiasm, cheer and meaningful discussions. I also want to
thank the Finkel lab members, who have always been so welcoming and never
minded me intruding in their lab activities. I cannot thank Yamini and Namita
iii
enough - they have been like sisters to me and have made my transition into a
completely new eld painless.
My special thanks to the Department of Chemistry at the University of South-
ern California for letting me be a part of the graduate program and providing me
with all the resources and help that I needed to make my way here.
I'm extremely grateful to my parents, Anuradha and Anand Karbelkar, for all
their support and encouragement. It was their dream that my sister and I should
receive the nest education and they worked hard, against all odds, to provide it
to us. I cannot imagine myself being here without their immense eorts. I also
want to thank my big sister Avanti, always the quiet shore by the rough seas, my
role model. I cannot begin to express my thanks to my aunt and uncle, Megha
and Dilip Bhumralkar, who have been there for me throughout these ve years in
so many ways that I have never missed home.
Above all, I would like to thank my best friend and companion, Chris. The
last few years have been a roller coaster ride but you have been my constant. You
have seen me through this journey and have always been so loving, helpful and
supportive. Meeting you was the best that happened to me. Thank you for helping
me keep it together, I owe you so much!
Last but not the least, I would like to acknowledge the funding agencies who
have made this research possible: This work was supported by the NASA Astrobiol-
ogy Institute (Life Underground project) under cooperative agreement NNA13AA92A,
and National Science Foundation Dimensions of Biodiversity Grant 1542527. Work
on human-associated microbes is supported by Air Force Oce of Scientic Research
Grant FA9550-14-1-0114.
iv
Contents
Dedication ii
Acknowledgments iii
List of Figures vii
Abstract xv
1 Introduction 1
2 Background 8
2.1 Cell Energetics and Oxidative Phosphorylation . . . . . . . . . . . . 8
2.2 Anaerobic Respiration and Dissimilatory Metal Reduction . . . . . 11
2.3 Extracellular Electron Transfer . . . . . . . . . . . . . . . . . . . . 12
2.4 Microbial Electrochemistry . . . . . . . . . . . . . . . . . . . . . . . 15
2.4.1 Bioelectrochemical Systems (BES) . . . . . . . . . . . . . . 16
2.4.2 Common Electrochemical Techniques . . . . . . . . . . . . . 17
2.4.3 Control Experiments . . . . . . . . . . . . . . . . . . . . . . 20
2.5 Atomic Force Microscopy . . . . . . . . . . . . . . . . . . . . . . . . 22
2.5.1 Tapping Mode Atomic Force Microscopy . . . . . . . . . . . 22
3 An Electrochemical Investigation of Interfacial Electron Uptake
by the Sulfur Oxidizing BacteriumThioclavaelectrotropha ElOx9 24
3.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24
3.2 Materials and Methods . . . . . . . . . . . . . . . . . . . . . . . . . 27
3.2.1 Cell Growth Conditions . . . . . . . . . . . . . . . . . . . . 27
3.2.2 Bioelectrochemical Measurements . . . . . . . . . . . . . . . 28
3.2.3 Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . 30
3.2.4 Protein Quantication . . . . . . . . . . . . . . . . . . . . . 30
3.2.5 Nitrate quantication . . . . . . . . . . . . . . . . . . . . . . 31
3.3 Results and Discussion . . . . . . . . . . . . . . . . . . . . . . . . . 31
3.4 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 40
v
4 Extracellular Electron Transfer in the Human Microbiome 42
4.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42
4.2 Materials and Methods . . . . . . . . . . . . . . . . . . . . . . . . . 47
4.2.1 Cell Growth Conditions . . . . . . . . . . . . . . . . . . . . 47
4.2.2 Bioelectrochemical Measurements . . . . . . . . . . . . . . . 48
4.2.3 Membrane Potential Analysis . . . . . . . . . . . . . . . . . 50
4.3 Results and Discussion . . . . . . . . . . . . . . . . . . . . . . . . . 51
4.3.1 Electrochemical investigation of electron shuttling mecha-
nism in Pseudomonas aeruginosa PA14 . . . . . . . . . . . . 51
4.3.2 Electrochemical enrichment and isolation of gut microbes
from human fecal samples . . . . . . . . . . . . . . . . . . . 62
4.4 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 66
5 Bacterial Immobilization for Atomic Force Microscopy under Phys-
iological Conditions 68
5.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 68
5.2 Materials and Methods . . . . . . . . . . . . . . . . . . . . . . . . . 71
5.2.1 Cell Growth Conditions . . . . . . . . . . . . . . . . . . . . 71
5.2.2 Substrate Preparation . . . . . . . . . . . . . . . . . . . . . 72
5.2.3 Sample Preparation . . . . . . . . . . . . . . . . . . . . . . . 73
5.2.4 Atomic Force Microscopy . . . . . . . . . . . . . . . . . . . . 73
5.2.5 Evaluation of cell viability . . . . . . . . . . . . . . . . . . . 74
5.2.6 AFM Image Processing . . . . . . . . . . . . . . . . . . . . . 74
5.3 Results and Discussion . . . . . . . . . . . . . . . . . . . . . . . . . 74
5.3.1 Surface modication and analysis . . . . . . . . . . . . . . . 76
5.3.2 Assessment of structure, size and viability of immobilized cells 77
5.3.3 Phase Image Analysis . . . . . . . . . . . . . . . . . . . . . 82
5.4 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85
6 Conclusion 87
Bibliography 90
vi
List of Figures
2.1 Representation of an electron transport chain of respiratory organ-
isms. Inner membrane bound redox proteins catalyze electron trans-
fer reactions along the chain, accompanied by the formation of a
proton gradient across the membrane. This proton gradient drives
the synthesis of adenosine triphosphate. Figure obtained from [80] . 9
2.2 Redox tower displaying electron donors and acceptors commonly
used by microorganisms. Chemicals are arranged according to their
standard reduction potentials at pH 7. More reducing species occupy
the top of the tower and serve as electron donors, whereas more oxi-
dizing compounds lie at the bottom. Red arrows classify microbes
based on their preferred donors, acceptors and metabolisms. . . . . 10
2.3 Representation of extracellular electron transfer in dissimilatory metal
reducing microbe Shewanella oneidensis MR-1. Electron transfer to
the outer membrane is facilitated by inner membrane and periplas-
mic redox proteins to outer membrane multiheme cytochromes. Out-
ward electron transfer to solid surfaces is enabled by direct con-
tact of cytochromes, cytochrome bound cofactors, membrane exten-
sions consisting of cytochromes or indirectly with the help of small
molecules. Figure obtained from [32] . . . . . . . . . . . . . . . . . 13
2.4 Proposed mechanisms of inward EET by microbes from a cathode.
a) Mediated electron transfer by cathodically generatedH
2
b) Medi-
ated electron transfer via H
2
generated by cell-secreted electrode-
attached hydrogenases c) Direct electron transfer using outer mem-
brane c-type cytochromes. Figure obtained from [96] . . . . . . . . 14
vii
2.5 Examples of three electrode bioelectrochemical systems (BES) used
to study extracellular electron transfer. a) Bioreactors used for most
bulk electrochemical analyses in this dissertation b) Gas-tight BES
used for monitoring gases in the reactor head space c) Bond lab
reactors [113] designed to provide better anaerobicity in the system
and enhanced control over mass transport properties d) Small BES
with optically transparent electrode based allowing electrochemical
measurements in conjugation with inverted microscopy. . . . . . . . 17
2.6 Cyclic Voltammetry (CV) of Geobacter sulfurreducens. a) Turnover
CV performed on an electrode-attached biolm (with acetate) at
1mV/s on graphite rod. Plot obtained from [113] b) Non-turnover
CV of starved biolm (no acetate) revealing redox peaks. Bot-
tom gure represents baseline subtracted non-turnover CV. Plot
obtained from [114] . . . . . . . . . . . . . . . . . . . . . . . . . . . 19
2.7 Representative chronoamperometry plot after media exchange. After
media exchange (arrow), if redox-active moieties are attached to
the electrode, the electrochemical activity is retained (green) and is
similar to before media exchange (black). A drop in current implies
redox molecules were present in the media and hence after exchange,
the electroactivity declines (dashed blue). . . . . . . . . . . . . . . . 21
2.8 Illustration of Tapping Mode Atomic Force Microscopy (AFM). Oscil-
lation amplitude of the tip (blue) changes on experiencing changes
in the topography of the surface. . . . . . . . . . . . . . . . . . . . 23
3.1 Cathodic activity of Thioclava electrotropha ElOx9. Chronoamper-
ometry measurements show an increase in cathodic current after
inoculation of cells into a bioreactor containing a working electrode
poised at -278 mV vs SHE. No activity is detected from an uninoc-
ulated abiotic control (black). . . . . . . . . . . . . . . . . . . . . . 32
3.2 Eects of inhibition on cathodic activity. a) Chronoamperometry of
Thioclava electrotropha ElOx9, with a working electrode poised at
-278 mV vs SHE, shows cathodic activity and subsequent collapse of
this activity after inhibition with 5 mM azide. b) Summary (n=8) of
cathodic currents detected from abiotic (sterile) electrodes, maximal
current after addition of T. electrotropha ElOx9 cells, and as a result
of inhibition with 5 mM azide. . . . . . . . . . . . . . . . . . . . . . 33
viii
3.3 Cathodic activity of Thioclava electrotropha ElOx9 without carbon
source. Chronoamperometry of T. electrotropha ElOx9 cells in the
absence of any inorganic carbon source (no bicarbonate in SWB
reactor media, and no CO
2
in purging gas). On inoculation of cells
in the bioreactor, with carbon cloth poised at -278 mV vs SHE,
cathodic current increases to similar levels as seen in experiments
with inorganic carbon sources. . . . . . . . . . . . . . . . . . . . . . 35
3.4 Turnover Cyclic Voltammetry (CV) of Thioclava electrotropha ElOx9.
Turnover CV (in presence of nitrate) of T. electrotropha ElOx9
reveals a catalytic wave indicating sustained extracellular electron
uptake from electrodes coupled to nitrate reduction by the cells.
The catalytic wave is retained after exchanging old media with fresh
media (purple). CVs of cell-free spent media shows no detectable
redox features. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 36
3.5 First derivative of turnover cyclic voltammetry (CV). a) Representa-
tive derivative CV from both forward and reverse scans. The width
at half-max is 105 mV. b) Derivative CVs of multiple experiments
(with varying cathodic activity) showing similar location and width
at half-max of 110 8 mV. Only reductive scan derivatives are
shown in (b) for clarity. . . . . . . . . . . . . . . . . . . . . . . . . 37
3.6 Microscopic characterization of electrodes. a) Scanning electron
microscopy of carbon cloth electrode bers after electrochemical
measurements of Thioclava electrotropha ElOx9 cells (Scale bar 10
m). Inset shows higher magnication of the same electrode and
attached cells (Scale bar 2 m). b) Fluorescence microscopy image
of the carbon cloth electrode bers shows attached cells using FM
4-64FX to stain cell membranes (Scale bar 10 m). . . . . . . . . . 38
3.7 Comparing turnover and non-turnover voltammetry. Turnover cyclic
voltammetry (CV) of electrode-attached Thioclava electrotropha ElOx9
cells (red) compared to an abiotic control (black). Removal of
nitrate results in a non-turnover CV (purple) revealing redox peaks
indicated by arrows. Catalytic activity is re-gained by the cells upon
re-introduction of nitrate (blue). . . . . . . . . . . . . . . . . . . . . 39
3.8 Eect of hydrogen addition. Turnover cyclic voltammetry of electrode-
attached Thioclava electrotropha ElOx9 cells before (red) and after
(blue) purging with 80% H
2
=CO
2
for 1 hour (blue). . . . . . . . . . 40
ix
4.1 Electrochemical analysis of Pseudomonas aeruginosa PA14 wild-
type (WT). a) Chronoamperometry measurements indicate an increase
in anodic current upon inoculation of P. aeruginosa PA14 WT cells
in a bioreactor containing a working electrode poised at 622 mV
vs SHE. The current remains steady over 50 hours indicating no
detectable amount of phenazines building up over time. b) Cyclic
voltammetry (CV) after 50 hours reveal no signicant change in
electrochemical activity when compared to abiotic control (black). . 52
4.2 Electrochemical analysis of Pseudomonas aeruginosa PA14phz. a)
Chronoamperometry measurements indicate an increase in anodic
current upon inoculation of P. aeruginosa PA14phz cells in a
bioreactor containing a working electrode poised at 622 mV vs SHE
with 20M pyocyanin (blue). No activity was detected in an uninoc-
ulated abiotic control containing only pyocyanin (black) or inocu-
lated control containing only cells (purple). b) Cyclic voltammetry
(CV) of pyocyanin indicates a reversible peak with mid-point poten-
tial at -21 mV (black). No activity is observed without pyocyanin
(purple). Suppressed current density with cells and pyocyanin (blue)
after 90 hours possibly occurs due to electrode passivation. . . . . . 54
4.3 Eect of glucose and nitrate on the anodic activity of Pseudomonas
aeruginosa PA14phz. a) Comparable anodic current densities
detected from cells in bioreactors with 20 mM electron donor glucose
(blue) and without glucose (purple). b) Addition of 25 mM nitrate
in order to substitute for the poised working electrode as an electron
acceptor for the cells, led to a rapid decline in anodic current (red)
as compared to the bioreactor without any nitrate (blue). . . . . . . 55
4.4 Anodic activity of cell-free inoculum of Pseudomonas aeruginosa
PA14phz. Chronoamperometry of cell-free inoculum of P. aerugi-
nosa PA14phz indicates no increase in anodic current upon inoc-
ulation in a bioreactor containing an electrode poised at 622 mV vs
SHE (red) compared to the abiotic control (black). . . . . . . . . . 55
x
4.5 Analysis of planktonic cell density. Summary (n=3) of bioreactor
planktonic cell density of P. aeruginosa PA14phz, expressed in
colony forming units per ml (CFU/ml), compared between Day 1
(day of bioreactor inoculation, dark gray) and Day 7 (light gray),
under dierent conditions { with cells, 20 M pyocyanin, 20 mM
glucose and working electrode poised at 622 mV vs SHE), with-
out poised working electrode, without added pyocyanin (PYO) and
without added glucose. A slight loss in planktonic cell density is
observed in all conditions but the rst. . . . . . . . . . . . . . . . . 56
4.6 Eects of stirring on the eciency of pyocyanin cycling. a) Chronoam-
perometry of P. aeruginosa PA14phz shows a dependence of anodic
current density on the extent of mixing in the bioreactor. The cur-
rent density is lowest in static bioreactors (red), higher in actively
purged bioreactors (blue) and highest in actively purged and stirred
bioreactors. b) The frequency of pyocyanin redox cycles between
P. aeruginosa PA14phz cells and the working electrode increased
with enhanced mixing of planktonic culture in the bioreactor. At 77
hours of chronoamperometry, the number of cycles calculated for a
static bioreactor increased from an average (n=3) of 5 cycles to 13
(with N
2
purging) and to 29 (with purging and stirring). . . . . . . 58
4.7 Experimental set-up for the analysis of cellular membrane potential.
a) Schematic representation of the bioreactor- cylindrical glass body
with optically transparent patterned gold coverslip at the base as a
working electrode, reactor top with counter and reference electrode.
Cells settled on the working electrode are stained from Thio
avin T

uorescent dye in reactor media and observed through an inverted
microscope. b) Patterned gold electrode used for the experiment -
glass coverslip coated with 5nm titanium and 10 nm gold. . . . . . 59
xi
4.8 Eect of applied potential on the membrane potential of P. aerug-
inosa PA14phz. a) Cells under aerobic conditions display high
Thio
avin T (ThT)
uorescence intensity before addition of 100
M carbonyl cyanide m-chlorophenyl hydrazine (CCCP) (scale bar 5
m). b) Cells under aerobic conditions display reduced Thio
avin T
(ThT)
uorescence intensity after addition of 100 M CCCP (scale
bar 5 m). c) ThT
uorescence intensity of cells increase over time
under aerobic condition followed by a sudden decrease after the addi-
tion of CCCP. d) No increase in ThT
uorescence (green) or current
(orange) was observed over time with a constant applied potential
of 622 mV vs SHE (purple). e) Comparable ThT
uorescence of
cells on the poised electrode (green) and on glass (black) indicates
no signicant gain in membrane potential for cells from the applied
potential. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61
4.9 Electrochemical activity of metacommunity human fecal sample.
Chronomaperometry measurements indicate an increase in anodic
current upon inoculation of the fecal community in a bioreactor with
a working electrode poised at 422 mV vs SHE (red). Addition of 10
mM cyanide, a respiratory chain inhibitor, led to a rapid decrease
in the anodic current (gray). No activity was observed in the sterile
abiotic control (black). . . . . . . . . . . . . . . . . . . . . . . . . . 62
4.10 Anodic activity of lysed and heat killed samples. a) Chronoamper-
ometry of fecal community samples either lysed (blue) or heat-killed
(purple) before inoculation in a bioreactor with working electrode
poised at 422 mV vs SHE produced no activity compared to reg-
ular inoculum. Lysed community control showed an in increase in
current after 45 hours. b) Chronoamperometry of all three colonies
isolated from lysed community control led to an increase in anodic
current upon inoculation in bioreactors poised at 422 mV vs SHE
(red, blue, purple). No activity was detected in the sterile abiotic
control (black). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63
4.11 Anodic activity of isolated colonies. Chronoamperometry of three
colonies, Enterococcus faecalis (red), Escherichia coli (blue) and
Klebsiella pneumoniae (purple), isolated from electrode attached
biolm of fecal community indicated an increase in anodic current
upon inoculation in a bioreactor containing a working electrode
poised at 422 mV vs SHE. . . . . . . . . . . . . . . . . . . . . . . . 65
xii
5.1 Illustration of phase response of an AFM tapping tip on a sample
with dierent material properties. As the tip is scanned across a
sample, the phase of the oscillating tip (red) relative to the driver
signal (black) changes when on the surface compared to when on
the cells. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 72
5.2 Schematic representation of immobilization technique. a) Silicon
wafer coated with Poly-L-lysine (PLL) with exposed amines. b)
Condensation reaction between exposed amines and glutaraldehyde
leads to the formation of imine bond. c) Amines on cell surface
of microbes undergo another condensation reaction with glutaralde-
hyde leading to crosslinks and cellular immobilization. . . . . . . . . 77
5.3 AFM analysis of silicon substrate. a) AFM image of uncoated silicon
wafer (Scale bar 100 nm). b) AFM image of silicon wafer coated
with Poly-L-lysine (PLL) and glutaraldehyde (Scale bar 100 nm). c)
Height prole of uncoated silicon wafer. d) Height prole of silicon
wafer coated with PLL and glutaraldehyde. . . . . . . . . . . . . . 78
5.4 Comparison of AFM images of cells under dry and under liquid con-
ditions. Tapping AFM amplitude images of a) Shewanella oneiden-
sis MR-1 b) Thioclava electrotropha ElOx9 c) Pseudomonas aerug-
inosa PA14 on silicon wafer under dry conditions. (Scale bar 4
m). Tapping AFM amplitude images of d) S. oneidensis MR-1
e) T. electrotropha ElOx9 F) P. aeruginosa PA14 cells immobilized
on silicon wafer and imaged under phosphate buered saline under
physiological conditions (Scale bar 1 m). . . . . . . . . . . . . . . 79
5.5 Height analysis of cells under liquid conditions. AFM height images
and corresponding line proles of a) Shewanella oneidensis MR-1
b) Thioclava electrotropha ElOx9 and c) Pseudomonas aeruginosa
PA14 displaying height across the drawn white line. (Scale bar 1m). 80
5.6 Viability assessment of immobilized cells. LIVE/DEAD staining of
a) Shewanella oneidensis MR-1 b) Pseudomonas aeruginosa PA14
and c) Thioclava electrotropha ElOx9 indicating membrane integrity
of the cells (Scale bar 5 m). . . . . . . . . . . . . . . . . . . . . . . 81
5.7 Qualitative assessment of adhesion. Phase contrast images with dis-
crete color scheme representing cell adhesion of a) Shewanella onei-
densis MR-1, b) Thioclava electrotropha ElOx9, c) Pseudomonas
aeruginosa PA14 (Scale bar 1 m) . . . . . . . . . . . . . . . . . . 83
xiii
5.8 Extent of cell adhesion. AFM phase images and corresponding line
proles of a) Shewanella onedensis MR-1 b) Thioclava electrotropha
ElOx9 and c) Pseudomonas aeruginosa PA14 displaying shift in
phase across the drawn white line (Scale bar 1m). d) Mean phase
shifts over several cells observed in S. oneidensis MR-1, T. elec-
trotropha ElOx9 and P. aeruginosa PA14. . . . . . . . . . . . . . . 84
xiv
Abstract
Extracellular electron transfer (EET) is a metabolic process that allows microor-
ganisms to oxidize or reduce solid electron donors or acceptors including insoluble
minerals. This phenomenon governs many biogeochemical cycles of major ele-
ments including iron, sulfur, manganese, carbon and nitrogen. This process can
be mimicked on favorably poised electrode surfaces and has important implications
in microbial electrochemical technologies including microbial fuel cells, wastewater
treatment, bioremediation and microbial electrosynthesis. The eciency of these
operations relies largely on our understanding of the pathways that dictate EET.
With biophysical, biochemical and genetic toolkit at our disposal, our under-
standing of EET mainly arises from studies performed on model metal reduc-
ing microbes belonging to the genera, Shewanella and Geobacter. Nonetheless,
the EET phenomenon is diverse and widespread and many of its aspects are
still unclear. (1) Outward EET mechanisms (cell-to-electrode) have been exhaus-
tively characterized, however, the reverse process of inward EET (electrode-to-cell)
remains poorly understood, predominantly due to the lack of model organisms and
diculty in laboratory handling of current model microbes. (2) The studies used
to explore the scope of EET have been restricted mostly to natural environments,
however, anoxic niches of the human body that harbor a vast diversity of microbes
may also house novel electroactive microbes that are still uncharacterized. (3)
xv
Techniques used for characterizing EET, especially bulk electrochemistry, reports
the behavior of a large microbial population but fails to reveal single-cell electron
transfer properties or even dierentiate between electroactive and non-electroactive
cells.
This thesis addresses these issues in three parts. In the rst part, we investigate
the possible electron uptake mechanisms of a recently isolated marine microbe,
Thioclava electrotropha ElOx9. This sulfur oxidizer has been previously shown
to respire oxygen using reduced electrodes as a stable carbon source. However, a
detailed electrochemical characterization of inward EET pathways is lacking and
only limited to aerobic conditions. Here, we use amperometric and voltammetric
tools to demonstrate that ElOx9 can couple oxidation of cathode to reduction
of nitrate when it is used the sole electron acceptor. Our data reveals that the
electron uptake by ElOx9 is facilitated by direct contact, using a redox center with
a formal potential of -94 mV vs SHE, instead of soluble redox mediators.
In the second part, we examine EET in host associated bacteria residing in
anaerobic regions of human lungs and gut. Pseudomonas aeruginosa PA14, a lung
microbe, has been previously shown to respire anodes using phenazines as a solu-
ble redox mediator. However, bioelectrochemical data endorsing this mechanism
have been limited. Here, we undertake a thorough electrochemical characteriza-
tion of PA14 phenazine mutant (phz) cells with exogenously added phenazines,
thereby conrming that phenazines enable electron shuttling between the cells and
an external electrode. We also demonstrate that the eciency of phenazine redox
cycling directly depends upon the extent of mixing in the bioelectrochemical sys-
tem. Additionally, we attempt to qualitatively determine whether PA14 (phz)
cells respiring anodes experience any bioenergetic gain, by conjugating amperomet-
ric analysis with
uorescence microscopy and using a
uorescent dye Thio
avin T,
xvi
capable of reporting changes in cellular membrane potential. Our results indicate
that the cells likely experience a low energetic gain due to low electron transfer
rates.
Pertaining to the anaerobic gut microbiome, most resident microbes are known
to undergo low energy yielding fermentation. However, other forms of metabolism,
including EET, may also be prevalent and remain unexplored. Here, we perform
anodic enrichment of facultative microbes from a community outgrown from human
fecal samples, representative of the large intestine. Seven morphologically distinct
colonies were isolated from electrode attached biolms, three of which were identi-
ed by 16S rRNA sequencing as Enterococcus faecalis, Klebsiella pneumoniae and
Escherichia coli. An increase in anodic current upon inoculation of these isolates
in bioreactors indicate possible EET capability of these microbes and are currently
under investigation.
In the third part, we focus on determining an ecient and consistent bacterial
immobilization technique applicable over diverse range of microbes for atomic force
microcopy (AFM) imaging under liquid. Here, we successfully immobilized three
phylogenetically diverse electroactive bacteria focused in this dissertation { She-
wanella oneidensis MR-1, Thioclava electrotropha ElOx9 and Pseudomonas aerug-
inosa PA14, on poly-L-lysine functionalized silicon wafers using 5% glutaraldehyde
as a crosslinker. Topographical images of the immobilized cells obtained from tap-
ping mode AFM revealed smooth surface structure in liquid as compared to dry-
ing related artifacts observed when imaged in air. With the help of
uorescence
microscopy using LIVE/DEAD stain, it was observed that most of the immobi-
lized microbes were viable and not aected by the chemical modication. Phase
contrast between the substrate and cells, obtained from phase images, was taken
xvii
as a measure of cell adhesion and its quantication revealed that ElOx9 cells were
strongly adhesive, followed by MR-1, then PA14 cells.
The work conducted in this thesis has important implications in understanding
the functional diversity of EET and the extent of environments where EET-capable
microbes may reside. T. electrotropha ElOx9, in addition to serving as a model
organism for understanding inward EET mechanism, may also help in understand-
ing biogeochemical sulfur oxidation in marine environments. A thorough mecha-
nistic characterization of gut microbes isolated as a result of electrode enrichment,
may further our understanding of unique anaerobic metabolisms in the gut envi-
ronment that had not been explored yet. Manipulation of EET pathways through
biosynthetic techniques can help externally control bacterial population associated
with human gut. Additionally, developing sophisticated tools like AFM-SECM
will enable detection of electroactive bacteria and simultaneous quantication of
their activity from a mixed culture of microbes.
xviii
Chapter 1
Introduction
Electron transfer is an elementary and pervasive class of reactions that broadly
dictates most chemical and biological processes [63]. Life originated and sustained
from electron transfer reactions [105]. In the cells of respiratory organisms, the
energy required to sustain life is derived from the transfer of electrons from reduced
donors to oxidized terminal electron acceptors [69].
Respiratory organisms possess specialized machinery to carry out electron
transfer. Eukaryotic cells rely on mitochondria [69], while bacteria rely on inner
membrane bound components of the electron transport chains in for this process
[68]. During the process of respiration, the proteins of electron transport chain
oxidize the end products of metabolic cycles and transfer electrons along the chain
to a terminal electron acceptor like molecular oxygen. This process is accompa-
nied by vectorial translocation of protons across the inner membrane [69]. Like
the hydrodynamic energy in dams, proton translocation causes an electrochemical
gradient to build up across the cell membrane driving the formation of adeno-
sine triphosphate (ATP), which is further used by cells for many enzymatic and
non-enzymatic functions [68, 69].
Aerobic organisms are restricted to using molecular oxygen as a terminal elec-
tron acceptor to produce ATP. Microbes, however, may possess dierent electron
transport chains allowing them to utilize alternate electron acceptors in the absence
of oxygen to assist the
ow of electrons [6]. Due to this metabolic
exibility, based
upon their habitats, microbes can use a variety of electron donors including organic
1
compounds, inorganic ions, sunlight or H
2
and electron acceptors such as nitrate,
sulfate, carbon dioxide or oxidized minerals to support electron
ow and ATP syn-
thesis. In addition to soluble electron donors and acceptors, microbes have also
evolved to respire insoluble minerals by a process termed as dissimilatory metal
reduction. Evidence suggests that this form of microbial metabolism is ancient and
helped shape the early Earth when minerals were abundant, and oxygen was lim-
ited [106]. However, microbial metal reduction has largely been ignored as a form of
anaerobic respiration until three decades ago when two microbes- Shewanella onei-
densis MR-1 [127] and Geobacter sulfurreducens [107] were isolated as manganese
and iron reducers, respectively. These microbes, also classied as dissimilatory
metal reducing bacteria (DMRB), were found to be capable of coupling mineral
reduction to energy conservation [10]. This form of metabolism where microbes
respire external abiotic minerals by transferring electrons across an insulating cell
membrane is called extracellular electron transfer (EET) [48].
The EET capable microbes present in the Earth's lithosphere and hydrosphere,
play an important role in governing the biogeochemical cycles of major elements
including iron, sulfur, manganese, nitrogen and carbon [83]. Microbes can also
mimic EET on favorably poised solid electrodes when they are used as proxies for
minerals. This opens avenues for understanding bioelectrochemical technologies
such as microbial fuel cells [160] or wastewater treatment processes [50], where
anode-respiring microbes can catalyze the breakdown of complex organic substrates
or waste matter and perform cell-to-electrode EET, thus generating electricity.
The success of these techniques largely depends upon our understanding of how
microbes interact with these abiotic surfaces for their metabolic gain. Redox active
protein conduits or small organic molecules that can carry electrons from metabolic
reducing equivalents from the microbe, assist in \wiring" it to a solid mineral or
2
electrode, enabling short-range or long-range EET. This broadly leads to the clas-
sication of EET mechanisms into direct contact or mediated electron transfer [32].
In direct contact EET, interfacial electron transfer is possible through direct con-
tact of either cells with outer membrane multiheme cytochromes or cell-produced
conductive laments containing multiheme cytochromes, with an external surface
[24, 49, 147]. In mediated EET, redox active molecules like
avins [112, 171], humic
acids [133] or phenazines [174], cycle electrons between the cells and the surface.
Ideally, these molecules should be able to shuttle electrons multiple times [96].
To date, our knowledge of EET is limited to studies focused on two classes of
metal reducing organisms, Gammaproteobacteria Shewanella [127] and Deltapro-
teobacteria Geobacter [107]. Thorough electrochemical, biophysical, biochemical
and genetic characterization of these microbial types has yielded a better under-
standing of outward (or anodic) EET mechanisms where microbes respire an exter-
nal solid electron sink [159]. However, the reverse process of inward (or cathodic)
EET is also gaining a lot of interest, especially in the context of sustainable biotech-
nology [149]. In bioelectrochemical systems, microbes capable of using cathodes
as electron donors are being studied for their potential applications in wastewater
treatment, bioremediation and production of biofuels (microbial electrosynthesis)
[70, 74, 79]. The mechanisms of microbial electron uptake from solid electron
donors, however, remain poorly understood due to the lack of model organisms
that are easy to cultivate and that allow electrochemical characterization under
anaerobic conditions.
Several previous studies have focused on enriching and isolating microbes capa-
ble of cathode oxidation [18, 120, 138]. To seek representative chemolithotrophic
microbes capable of performing inward EET, one such study subjected marine sed-
iments of Catalina Harbor, CA to electrochemical enrichments on cathodes poised
3
at dierent reducing potentials mimicking those of reduced iron and sulfur minerals
native to the sediments [151]. This study led to the isolation of several sulfur and
iron oxidizers capable of using a cathode as a stable source of electron. In chap-
ter 3, we explore electrochemical properties of one of the sulfur-oxidizing isolates
obtained from this study, Thioclava electrotropha ElOx9 and evaluate its potential
in serving as a model organism to study inward EET.
Another aspect of EET that remains under-explored is the diversity of this
phenomenon across various environments. While the model organism, Shewanella
oneidensis MR-1 and Geobacter sulfurreducens, are both representatives of aquatic
environments, EET is widespread and has been reported in other environments
including the deep subsurface, marine sediments [151], ocean cold seeps [148],
soil, hydrothermal vents [184], acid mine drainage [17], etc. However, microbes
existing in niches beyond the environmental and engineered systems, for instance,
in anaerobic zones in the animal ecosystem may also perform EET. For example,
anaerobic sections of the human respiratory and digestive systems (large intestine)
may harbor microbes capable of respiring oxidants beyond molecular oxygen or
soluble inorganic ions, such as particulate matter or human epithelial cells.
To represent EET in anoxic respiratory airways, Pseudomonas aeruginosa
PA14 is a well-studied model lung bacterium that has been reported to rely on
endogenously produced N-heterocyclic compounds, phenazines, to serve as elec-
tron shuttles to mediate long-range EET with an oxidized electrode [174]. However,
the lack of electrochemical data exhibiting evidence for phenazine redox shuttling
casts ambiguity towards the said mediated EET pathway. On the gut microbiome
front, the dearth of electroactive microorganisms representing this environment,
hinders our understanding of EET metabolisms prevalent in the diverse ecosystem
of human large intestine. Hence, in chapter 4, we attempt at characterizing EET in
4
the human associated microbiome with a focus on lung and gut environments. We
undertook electrochemical approaches to thoroughly characterize EET in P. aerug-
inosa PA14. Additionally, in collaboration with the lab of Dr. Steven Finkel, we
take an exploratory approach to electrochemically enrich for microbes in human
fecal samples capable of respiring oxidized electrodes in pursuit of electroactive
isolates representative of the human gut ecosystem.
One of the most common techniques used for studying EET includes bulk anal-
ysis in bioelectrochemical systems where electrochemical response of a microbial
population with respect to a poised electrode is monitored. Hyphenating electro-
chemical studies with microscopy can help understand subpopulation or single-cell
dynamics of EET [66]. Scanning Electrochemical Microscopy with Atomic Force
Microscopy (AFM-SECM) is a powerful technique which when implemented on
immobilized cells can potentially quantify localized current densities and gener-
ate a map of cellular electroactivity in conjugation with structural information of
the microbe at nanoscale resolution [11]. On a multispecies bacterial platform,
AFM-SECM can assist in identication and dierentiation of electroactivity and
morphology of EET-capable microbes from the ones that cannot perform EET.
One important aspect of executing this technique is the requirement to perform
AFM under a liquid droplet. While physiologically more relevant, immobilization
of bacteria is one of the major problems for AFM imaging in liquid media as the
cells tend to detach easily due to the lateral forces of scanning tip [47]. Therefore, in
chapter 5, we investigate protocols used for bacterial immobilization to determine
an ecient technique that consistently allows for AFM imaging of electroactive
bacteria under physiologically relevant conditions. This will serve as the rst step
to enable exploration of more complex techniques such as AFM-SECM.
5
Overall, this thesis exploits electrochemical and imaging tools to capture the
extent and diversity of EET. This thesis contains 6 chapters. Chapter 2 elabo-
rates on cellular energetics and its relevance in aerobic and anaerobic respiration,
the meaning of extracellular electron transfer and the pathways of electron trans-
fer adopted by metal reducing and metal oxidizing microbes. This chapter also
includes a detailed introduction to microbial electrochemistry which is a popu-
lar tool used to evaluate extracellular electron transfer and an account of various
controls that are commonly performed to help deduce the mechanism of electron
transfer.
Chapter 3 describes a detailed electrochemical investigation to elucidate extra-
cellular electron uptake mechanism(s) in sulfur oxidizing T. electrotropha ElOx9
under anaerobic conditions. We observed that ElOx9 was capable of coupling cath-
ode oxidation to nitrate reduction. Cyclic voltammetry in the presence of nitrate
revealed a sigmoidal feature indicating multiple turnovers of redox proteins. Con-
trol experiments conrmed a direct contact-based electron transfer mechanism (no
soluble mediators or electrode attached enzymes involved). We were also able to
establish the potential window of electroactivity of ElOx9 cells using carbon cloth
as a stable electron source.
Chapter 4 aims to understand extracellular electron transfer in microbes asso-
ciated with the human microbiome. A comprehensive electrochemical analysis was
performed on model lung bacterium P. aeruginosa PA14 to bridge gaps between
previously existing studies. We were able to conrm that PA14 uses phenazines as
electron shuttles to mediate electrons between cells and external electrode and that
the eciency of redox shuttling depends upon the extent of agitation of bioreactor
environment. Additionally, we performed electrochemical enrichment of human
fecal samples and successfully isolated three facultative electroactive bacteria {
6
Enterococcus faecalis, Escherichia coli and Klebsiella pneumoniae, all representa-
tive of the human gut environment.
Chapter 5 highlights our attempts towards successfully immobilizing various
electroactive bacteria including S. oneidensis MR-1, T. electrotropha ElOx9 and
P. aeruginosa PA14, to allow atomic force microscopy under liquid. We concluded
that the use of a dialdehyde crosslinker (glutaraldehyde) consistently immobilized
cells on amine functionalized surface via covalent bonding without majorly aect-
ing their viability. We used phase images obtained from intermittent contact AFM
to determine the extent of adhesion and immobilization of the microbes, observ-
ing that the immobilization procedure worked most strongly for ElOx9 cells, then
MR-1 and PA14 cells.
Chapter 6 concludes this thesis in which I brie
y review the work done and pro-
vide an outlook into the future work that can be done to further our understanding
of extracellular electron transfer in poorly explored environments.
7
Chapter 2
Background
2.1 Cell Energetics and Oxidative Phosphoryla-
tion
Electron transfer reactions are central to all life processes. Essential cellular oper-
ations like respiration, fermentation and photosynthesis rely on redox reactions
driven by electron
ux from a reduced electron donor to an oxidized electron
acceptor [69]. Microbial periplasmic membrane harbors an electron transport chain
which is essential to carry out electron transduction and generation of an electro-
chemical gradient [68]. This gradient drives the synthesis of universal energy cur-
rency adenosine triphosphate (ATP) from adenosine diphosphate (ADP). Energy
released from the dissociation of ATP is utilized to carry out most chemical and
mechanical functions of a cell, and invariably utilized to drive all biology [68] (Fig-
ure 2.1).
Bacterial electron transport chain comprises of various electron transfer pro-
teins such as dehydrogenases, oxidoreductases and cytochromes, all equipped with
redox components: Fe-S clusters, hemes, quinones and
avins that enable electron

ow [93]. During cellular respiration under aerobic conditions, chemotrophic aer-
obes or facultative anaerobes may undergo oxidative phosphorylation. Reducing
equivalents in the form of NADH andFADH
2
generated at the end of glycolysis
in the cytoplasm, are oxidized by NADH dehydrogenase, where the electrons are
further passed on to the other protein complexes down the electron transport chain.
8
Figure 2.1: Representation of an electron transport chain of respiratory organ-
isms. Inner membrane bound redox proteins catalyze electron transfer reactions
along the chain, accompanied by the formation of a proton gradient across the
membrane. This proton gradient drives the synthesis of adenosine triphosphate.
Figure obtained from [80]
These complexes are wired together with the help of lipophilic soluble molecules,
quinones [93]. Flow of electron is governed by the dierence in redox potentials of
the components of the chain [111]. Final fate of electrons is the reduction of an
oxidized electron acceptor (e.g. oxygen in case of aerobic respiration). Many of
the reductases and dehydrogenases can also act as proton pumps, thus can cou-
ple electron transport with proton translocation across inner membranes, thereby
generating an electrochemical gradient called the proton motive force (PMF) [68].
The enzyme ATP synthase, also embedded in the inner membrane, is driven by
the PMF to drive phosphorylation (addition of phosphate ion) of ADP to form
ATP [111].
9
Figure 2.2: Redox tower displaying electron donors and acceptors commonly used
by microorganisms. Chemicals are arranged according to their standard reduction
potentials at pH 7. More reducing species occupy the top of the tower and serve as
electron donors, whereas more oxidizing compounds lie at the bottom. Red arrows
classify microbes based on their preferred donors, acceptors and metabolisms.
For their existence, survival and growth, cells must generate ATP which is
possible by electron transfer from an electron donor to acceptor. The electron
transfer is feasible if the donor has more negative reduction potential compared to
the acceptor, whereas, the drive for electron transfer depends upon the dierence
between reduction potentials of the donor and acceptor as shown in Figure 2.2
[111].
10
2.2 Anaerobic Respiration and Dissimilatory
Metal Reduction
Microbes exist in highly heterogeneous environments such as deep subsurface or
marine sediments and can extract free energy by driving electron transfer reac-
tions across redox gradients for their maintenance and survival. While oxygen is
the most energetically favorable electron acceptor, in these environments, such as
in an unperturbed marine sediment, oxygen supply is not only limited given its
low solubility in water, but also depletes rapidly with depth [130]. However, this
allows gradients to form in the deeper sediments with alternate electron acceptors
made available to the microbes, usually in the order decreasing redox potential
[130]. As a result, microbes use these electron acceptors as a substitute for oxy-
gen and engage in anaerobic respiration. Apart from soluble electron acceptors,
such as nitrates or sulfates, metal ions in the form of Fe, Mn or S minerals are
also present in abundance [130]. Some microorganisms have evolved to exploit
oxidized minerals as their terminal electron acceptors to respire and gain energy.
This process is called dissimilatory metal reduction and the microbes performing it
are called dissimilatory metal reducing bacteria (DMRB). An example of DMRB is
Shewanella oneidensis MR-1 which was isolated from Lake Oneida, NY at depths
with limited oxygen but plenty of oxidized Mn [127]. This microbe is shown to
be capable of reducing an extracellular electron acceptor (oxidized Mn) and cou-
ple it with respiration [128]. Metal ions can also act as energy source for some
microbes. Lithotrophic organisms can use solid minerals as electron donors for
autotrophic growth. Metal oxidizing bacteria withdraw extracellular electrons from
a reduced mineral and couple them with reduction of oxygen, nitrate or carbon
11
dioxide. Rhodopseudomonas palustris [20], Mariprofundus ferroxydans [163] and
Acidithiobacillus ferroxidans [27] are some examples of iron (II) oxidizing microbes.
2.3 Extracellular Electron Transfer
In the absence soluble electron donors and acceptors, electroactive bacteria have
evolved to utilize solid state abiotic surfaces to respire. This capability of microbes
to link internal cellular metabolic processes to an external solid surface by con-
duction of electrons across an insulating cell membrane, is called extracellular
electron transfer (EET) [159]. The pathways by which microbes can perform EET
are broadly classied into two categories { (1) direct EET via direct contact [24] or
conductive laments [49, 147] and (2) indirect EET using cell secreted small redox
molecules [26] (Figure 2.3). In outward electron transfer, where microbes respire
external surfaces as terminal electron acceptors (TEA), short range electron trans-
fer is accomplished by microbes making a direct contact with the TEA. DMRB,
such as S. oneidensis MR-1 and Geobacter sulfurreducens in which these pathways
have been most thoroughly characterized, consist of multiheme c-type cytochromes
situated on their outer membranes. In S. oneidensis MR-1, an electron channel is
created with the help of multiheme proteins spanning across the bacterial mem-
brane. CymA present in the periplasmic membrane (oxidizes quinols) and transfers
electrons to the outer membrane protein conduit { MtrA, MtrB and MtrC with
the help of FccA and STC. MtrC is exposed on the outer membrane of the bacteria
which comes in direct contact with solid surfaces [159]. Presence of multiheme c-
type cytochromes also dictate outward EET in G. sulfurreducens strains DL-1 and
PCA. Periplasmic membrane of this microbe harbors ImcH and CbcL, both pro-
teins known to oxidize quinols from quinol pool. Electrons are transferred onward
12
Figure 2.3: Representation of extracellular electron transfer in dissimilatory metal
reducing microbe Shewanella oneidensis MR-1. Electron transfer to the outer
membrane is facilitated by inner membrane and periplasmic redox proteins to outer
membrane multiheme cytochromes. Outward electron transfer to solid surfaces is
enabled by direct contact of cytochromes, cytochrome bound cofactors, membrane
extensions consisting of cytochromes or indirectly with the help of small molecules.
Figure obtained from [32]
.
13
Figure 2.4: Proposed mechanisms of inward EET by microbes from a cathode.
a) Mediated electron transfer by cathodically generated H
2
b) Mediated elec-
tron transfer via H
2
generated by cell-secreted electrode-attached hydrogenases
c) Direct electron transfer using outer membrane c-type cytochromes. Figure
obtained from [96]
to porin-cytochromes complexes situated in the outer membrane with the help of
PpcA and PpcD. Cytochromes OmcB, OmcC, OmaB and OmaC are embedded
in porins OmbB and OmbD, forming terminal electron transfer protein complex
of the cell before passing electrons to solid surfaces [159]. Long range electron
transfer is possible with the help of conductive laments that these microbes can
form. S. oneidensis MR-1 form periplasmic extensions with c-type cytochromes
assembled across the length [137, 162]. In G. sulfurreducens, polymers of hexaheme
cytochromes OmcS (previously thought as type IV pili) form conductive laments
called nanowires [172]. Electron transfer through conductive extensions can also be
categorized as direct EET. Indirect EET relies on electroactive microbes secreting
small molecules such as
avins [112, 171], humic acids, quinones [133] or phenazines
[175] that can mediate electrons between the bacteria and an external acceptor via
diusion.
Inward EET is the reverse process of outward EET, where microbes accept elec-
trons from reduced electrodes (cathode) or minerals and further reduce TEA such
14
as carbon dioxide, nitrate, sulfate or oxygen. Inward EET mechanisms are poorly
understood due to the lack of model organisms. While still unclear, currently pro-
posed mechanism of electron uptake by bacteria includes (1) direct electron transfer
through outer membrane proteins [31, 150, 152] (2) mediated electron transfer via
cathodically produced small molecules (e.g. H
2
) [8] or (3) mediated electron trans-
fer through small molecules generated by cell-secreted electrode-attached enzymes
[39]. Both S. oneidensis and G. sulfurreducens are capable of cathode oxidation
as well. Evidence of S. oneidensis using the same MtrABC pathway as it uses in
anode reduction has been provided previously [150, 152], while G. sulfurreducens is
suspected to use a dierent pathway [161]. Other bacteria such as iron (II) reducing
Acidithiobacillus ferroxidans [27], M. ferroxydans [163], R. palustris [20] and iron
(0) reducing or biocorrosive Desulfovibrio ferrophilus [38] and Desulfopila corrodens
[15] have also been shown to use reduced electrodes as an electron source. Carbon
dioxide reducing microbes, especially acetogens and methanogens have also been
studied as important biocatalysts for microbial electrosynthesis [31, 132]. While
mechanistic studies are impending, metagenomic and metatranscriptomic studies
have implicated a porin-cytochrome complex Cyc2 to be functioning as an iron
(II) oxidase in the many iron (II) oxidizing bacteria during EET [28, 54].
2.4 Microbial Electrochemistry
The ability of electroactive microbes to transfer electrons across their outer mem-
branes and interact with external poised electrodes allows for the quantication of
electron
ux, electron transfer kinetics, recognition of rate limiting steps and iden-
tication of EET pathway(s) involved [97]. Electroactive bacteria (EAB) interact
with external electrodes either anodically (transferring electrons to electrode) or
15
cathodically (accepting electrons from electrode). Various factors aect the elec-
troactivity of electrode attached biolms, including but not limited to electrode
material and surface area, electrode potential, pH, presence or absence of oxygen,
types of electron donors and carbon source and presence of redox shuttles [16].
Due to the complex nature of the biolms formed on electrodes displaying both
bulk and local chemical and electrochemical gradients, the overall change in the
bioelectrochemical system is used an argument to explain changes in the electro-
chemical signals. However, important control experiments must be performed to
identify causes of variation of microbial electroactivity over time, which will be
discussed later in this chapter.
2.4.1 Bioelectrochemical Systems (BES)
To study microbial electrochemistry, single-chambered three electrode bioreactors
are used as BES [85]. In this conguration, a working electrode, a counter elec-
trode and a reference electrode are placed in a common electrolyte (minimal media)
without any physical separation of the anode or cathode [16]. The working elec-
trode is poised at a stable potential with respect to the reference electrode with
a known potential (generally, Ag/AgCl or calomel electrodes). Electroactivity of
the biolm is monitored at the working electrode. Counter electrode acts as an
auxiliary electrode that directs the
ow of electrons to or from the working elec-
trode, thereby completing the circuit. Maintenance of the potential dierence and
monitoring the current generated at the working electrode is possible with the help
of a potentiostat [11]. The bioreactors must be equipped with gas exchange ports
in order to control the BES environment. To attain anaerobicity, bioreactors are
purged with inert gases e.g., N
2
or other gas mixtures e.g., N
2
=CO
2
or H
2
=CO
2
depending upon the metabolic needs of the microbe. Electrode materials used in
16
Figure 2.5: Examples of three electrode bioelectrochemical systems (BES) used to
study extracellular electron transfer. a) Bioreactors used for most bulk electro-
chemical analyses in this dissertation b) Gas-tight BES used for monitoring gases
in the reactor head space c) Bond lab reactors [113] designed to provide better
anaerobicity in the system and enhanced control over mass transport properties
d) Small BES with optically transparent electrode based allowing electrochemical
measurements in conjugation with inverted microscopy.
BES should be cheap, biocompatible, inert and highly conductive. Commonly used
electrodes are carbon ber, carbon paper, graphite rod or carbon brushes [85]. A
few instances of commonly used three electrode bioreactors are shown in Figure
2.5
2.4.2 Common Electrochemical Techniques
Chronoamperometry
Chronoamperometry is one of the most common tools of electrochemical analysis
used to study microbial metabolism. In this experiment, a working electrode is
17
polarized at a constant potential over a certain time, which is used by microbes to
respire or ferment, consequently leading to an increase in current over the back-
ground (non-faradaic) current. If the electroactive components of a microbe are
not known, the microbes are allowed to get acclimatized to the electrode at open
circuit and polarization potential is chosen by gradually changing the potential of
the electrode attached biolm and determining where current is produced (voltam-
metric approach) [97]. These experiments determine whether a sustainable current
can be produced by the bacteria over extended periods of time [16].
Cyclic Voltammetry
Following controlled potential experiments, when a faradaic current is detected,
cyclic voltammetry (CV) helps in identifying the electroactive sites that may be
leading to an increase in current. In this experiment, potential is scanned at a
certain rate from initial to nal potential, back to the initial potential. As a result,
redox couples active within the tested range appear as reduction and oxidation
current peaks in the cycle [16]. CV is an important tool to identify the mechanisms
of EET { whether soluble redox shuttles are involved or whether electrode attached
redox sites are responsible for electron transfer { thereby distinguishing between
mediated and direct electron transfer mechanisms. Care must be taken while
selecting a potential window as to not harm the cells by strong potentials. CV is
performed either in the presence or absence of substrates, both reveling important
information about microbial metabolism and electron transfer.
Cyclic Voltammetry in the presence of substrate (Turnover CV): Under turnover
conditions, the soluble substrate is provided to the cells in excess and its diusion
to the electrode is not limiting. Consider the case study of Geobacter sulfurre-
ducens biolms from [113]. In turnover conditions, acetate is provided in excess.
18
Figure 2.6: Cyclic Voltammetry (CV) of Geobacter sulfurreducens. a) Turnover CV
performed on an electrode-attached biolm (with acetate) at 1mV/s on graphite
rod. Plot obtained from [113] b) Non-turnover CV of starved biolm (no acetate)
revealing redox peaks. Bottom gure represents baseline subtracted non-turnover
CV. Plot obtained from [114]
Scanning the applied potential gives rise to a sigmoidal wave form (Figure 2.6A).
The sudden increase in anodic current corresponds to electrons
owing continu-
ously from acetate to bacteria to the electrode. This is a zone of catalysis where
the electrons are produced by the cells faster than they can be transferred to the
electrode and hence are present in excess. In other words, the voltammogram is
limited by interfacial electrode kinetics. Further increasing the potential gives a
plateau which is representative enzymatic kinetics when it reaches saturation. It
is important to note that scan rates for microbial voltammetry must be kept slow
(1-5 mV/s) so that slow reactions have time to undergo completion and proteins
can turnover multiple times at a driving potential. To study catalytic properties
of proteins, it is important to ensure that diusion is not limiting, by keeping scan
rates low [97].
Cyclic Voltammetry in the absence of substrate (Non-Turnover CV): Under non-
turnover conditions, electrode attached biolm is starved by not providing any
19
substrate to the cells. CV performed under this condition at low scan rates, in the
same potential window as turnover CV, reveals the redox potentials of proteins
likely causing electroactivity of the cells. Proteins attached to the electrode give
rise reduction/oxidation peaks on scanning the potential and may not superimpose
like peaks from surface adsorbed redox moieties because of biolm heterogeneity
and protein orientation [114]. Non-turnover CV in G. surfurreducens is illustrated
in Figure 2.6B.
2.4.3 Control Experiments
Given the complex nature of electrode attached biolms, all bioelectrochemical
measurements must be performed against an abiotic control that is operated under
the same conditions, only without cells. Any changes observed in comparison to
the abiotic control can hence be attributed to the cells.
Media Exchange
This control is performed after a long-term electrode polarization experiment or
chronoamperometry and/or voltammetry when substantial faradaic current has
been observed. As the name suggests, old media from a BES is replaced with fresh
media. This control serves two purposes: (1) Identifying whether observed faradaic
reactions arise from electrode attached components such as cells or proteins. On
replacing media, if the electroactivity persists, it implies that surface adsorbed
moieties are indeed responsible for electron transfer [82, 114]. (2) to keep cells
viable in lengthy chronoamperometry experiments. Generally, in batch reactors,
current depletion is observed either due to cells running out of electron donor
or build-up of toxic by-products. Exchanging media in such cases, replenishes
20
Figure 2.7: Representative chronoamperometry plot after media exchange. After
media exchange (arrow), if redox-active moieties are attached to the electrode, the
electrochemical activity is retained (green) and is similar to before media exchange
(black). A drop in current implies redox molecules were present in the media and
hence after exchange, the electroactivity declines (dashed blue).
nutrients keeping cells active for longer duration, eects of which is observed by
an increase in current [76].
Spent Media
While media exchange control provides proof for direct electron transfer, spent
media control is performed to detect any soluble redox mediators present in the
reactor media possibly mediating electron transfer. This control is also performed
after some electrochemical activity is detected in the bioreactor. To prepare spent
media, cells from the bioreactor media are removed either by ltration or by con-
tinuously centrifuging and decanting the culture. If any electron mediators are
present in the spent media, CV performed in a fresh bioreactor with clean carbon
cloth will reveal diusion controlled redox peaks indicative of the mediator.
21
Addition of Inhibitors
Metabolic activity of cells in BES performing EET is conrmed by addition of
electron transport chain inhibitors. If the electron transfer pathway involved in
EET is known, inhibitors specic to that pathway can be added to conrm whether
there is inhibition of the pathway proteins, observed in the form of a decline in
current compared to uninhibited control. Dierent components of electron trans-
port chain may be directly involved with coupling cellular metabolic activity with
external electron transfer [29]. Given that each component is aected by a dierent
inhibitor, adding them separately, one at a time, can help in identifying crucial
electroactive components in the microbial electron transport chain.
2.5 Atomic Force Microscopy
Atomic Force Microscopy (AFM) is a scanning probe technique which relies on
weak interactions between a sharp tip and the sample, where the tip is scanned over
the sample and the interactions are controlled and quantied [12]. A piezoelectric
stage is used to precisely control and monitor the movements of sample stage and
tip. A feedback loop is used to maintain a constant height between sample and
tip during the raster scan. As a result, a topographic map of sample height with
respect to the lateral position of tip, is obtained. AFM is commonly used for the
study of biological samples because it allows sub-nanometer resolution without any
harsh sample preparation procedures [45].
2.5.1 Tapping Mode Atomic Force Microscopy
In tapping mode AFM, the cantilever gently taps (or intermittently contacts) the
surface in order to reveal topographical information. With the help of a piezo, the
22
Figure 2.8: Illustration of Tapping Mode Atomic Force Microscopy (AFM). Oscilla-
tion amplitude of the tip (blue) changes on experiencing changes in the topography
of the surface.
tip is made to oscillate vertically at its resonance frequency . Prior to engaging the
tip at the sample surface, the tip oscillates freely in air at its resonance frequency
at a constant amplitude. On bringing the tip closer to the surface, it experiences
attractive or repulsive forces, causing a shift in the resonant frequency and leading
to a drop in the oscillation amplitude . The change in oscillation amplitude is com-
pensated by movement of the tip back to the user-dened setpoint amplitude with
the help of a feedback loop. This mechanism generates a three-dimensional map
of the sample topography [125]. Tapping mode AFM circumvents the problems
associated with more commonly used contact mode AFM in which lateral forces
can damage soft biological samples [144].
23
Chapter 3
An Electrochemical Investigation
of Interfacial Electron Uptake by
the Sulfur Oxidizing Bacterium
Thioclava electrotropha ElOx9
3.1 Introduction
Microbial electrochemical systems (MES) that convert chemical energy to electri-
cal energy or vice versa are catalyzed by sustained microbial activity at electrode
interfaces [9, 149]. Central to this process is the respiratory strategy known as
extracellular electron transfer (EET), which can extend the intracellular electron
transport chains of microbes to external electrodes. In addition to playing a cru-
cial role in MES, microbes capable of EET drive major elemental cycles in natural
environments [53, 131], and can be harnessed for wastewater treatment [187], as
well as bioremediation of toxic metals [123]. Fullling the promise of these appli-
cations hinges on a better understanding of the fundamental EET mechanisms at
the microbe-electrode interfaces.
Outward EET allows microbes to respire external solid electron acceptors rang-
ing from minerals to anodes and has been extensively studied [62, 158, 159],
24
especially in the dissimilatory metal reducing bacteria Shewanella and Geobac-
ter [107, 127]. The established pathways of outward EET include (1) direct
contact-based electron transfer through outer membrane multiheme cytochromes
or nanowires [24, 49, 147], or (2) mediated electron transfer via soluble redox
shuttles such as
avins [112, 171]. Such mechanisms underlie the operation of
microbial fuel cells where microbes are harnessed to couple the oxidation of fuels
to electricity generation at anodes. As interest in the reverse process of microbial
electrosynthesis (electricity-to-fuel) continues to grow, the last few years have wit-
nessed signicant interest in the characterization of inward EET from cathodes to
microbes that reduce CO
2
and form biofuels [132, 149, 167]. However, our under-
standing of the molecular basis and cathode-microbe electron transfer mechanisms
remains limited compared to microbe-anode interactions. Current hurdles include
the lack of extensive characterization in model organisms (in the same vein as
Shewanella and Geobacter on anodes), slow growth rates, and general diculty in
culturing and genetically manipulating the autotrophic mineral oxidizing species
likely to be good candidates for cathodic electron uptake.
Despite these hurdles, recent studies demonstrated interfacial electron uptake
from cathodes by a number of microbes. Both model anode respiring bacteria, She-
wanella oneidensis MR-1 and Geobacter sulfurreducens, are capable of bidirectional
EET. In S. oneidensis, the Mtr multiheme cytochrome conduit involved in anodic
EET is also thought to be involved in electron uptake from cathodes [150, 152].
Recently, the environmental isolates Alcaligenes faecalis [173] and Ardenticatena
maritima [88] have also shown inward EET functionality. Iron oxidizing bacte-
ria, including Acidithiobacillus ferroxidans [27], Mariprofundus ferrooxydans [163],
and Rhodopseudomonas palustris [20] can also use electrodes as stable electron
sources. Interest in microbial electrosynthesis has also motivated studies where
25
methanogens and acetogens couple electron transfer from cathodes to CO
2
reduc-
tion, leading to methane or acetate production [8, 31, 39, 132]. Electrochemical
analyses of sulfate reducing bacteria involved in iron oxidation and steel corro-
sion, including Desulfopila corrodens [15] and Desulfovibrio ferrophilus [38], sug-
gest direct electron uptake mechanisms. The sulfur oxidizing bacteria Thiobacil-
lus denitricans and Sulfurimonas denitricans also exhibited signicant cathodic
activity [37, 185]. Surveying the aforementioned electrochemical studies, the pro-
posed models for inward EET can be broadly categorized as resulting from (1)
direct contact via redox proteins; (2) mediated electron uptake via small molecules
includingH
2
; and (3) through mediators (e.g. H
2
) generated by cell-produced but
electrode-attached enzymes (e.g. hydrogenases). However, detailed mechanisms
are still lacking, particularly when it comes to the precise identity and operat-
ing potentials of the putative charge carriers responsible for direct contact inward
EET.
Our understanding of inward EET will benet from increased eorts to isolate
and electrochemically characterize new model microbes, both to assess the existing
models and unravel novel mechanisms. In a study aimed at increasing the num-
ber of candidate microbes capable of inward EET, electrodes poised at reducing
potentials were previously used to enrich for chemolithotrophs from the marine
sediments of Catalina Harbor, CA [151]. A series of electrochemical (various set
potentials) and chemical (various oxidized Fe and S substrates) enrichments led to
the isolation of multiple new strains. One of the strains, Thioclava electrotropha
ElOx9, was isolated as a novel sulfur oxidizer from an enrichment with insoluble
elemental sulfur S
0
as the electron donor and nitrate as the electron acceptor. T.
electrotropha ElOx9 is a facultatively anaerobic alphaproteobacterium that can
grow using oxygen or nitrate as electron acceptors. In addition, T. electrotropha
26
ElOx9 can grow autotrophically with H
2
or reduced sulfur (S
2
, S
0
or S
2
O
2
3
)
as electron donors, or heterotrophically on organic substrates such as glucose and
acetate [30].
The metabolic
exibility of T. electrotropha ElOx9 makes it a promising organ-
ism for inward EET studies and, indeed, a previous report provided evidence that
it can couple electron uptake from a cathode toO
2
reduction[30]. However, inward
EET under anaerobic conditions has not been previously demonstrated and the
broad underlying mechanism (e.g. direct electron transfer, electron shuttling inter-
mediates, role of secreted extracellular enzymes) remains unclear. Since a detailed
electrochemical characterization of the microbial contribution to electron uptake
can be hindered by the inherent electroactivity of O
2
under the reducing poten-
tials used, measurements under anaerobic conditions will help disentangle biotic
and abiotic contributions to the observed cathodic activity. Here, we demon-
strate and characterize cathodic electron uptake by T. electrotropha ElOx9 under
nitrate-reducing conditions and propose that inward EET is facilitated by a direct
(contact-based) EET mechanism, rather than soluble intermediate electron carri-
ers. Our study suggests that strain ElOx9 is a highly promising model system for
cathode-microbe interactions and motivates future physiological and biochemical
studies of the precise molecular pathway through which it acquires electrons from
solid surfaces.
3.2 Materials and Methods
3.2.1 Cell Growth Conditions
Thioclava electrotropha ElOx9 was grown aerobically in Difco
TM
Marine (DM)
broth from a frozen stock (80
o
C in 30% glycerol), at 30
o
C and 200 rpm for 24
27
hours till stationary phase (7.4 x 10
9
CFU/ml) was attained. The overnight culture
was used to inoculate 100 ml of dened Salt Water Base (SWB) media at 2% (v/v)
and grown anaerobically for another 24 hours at 30
o
C and 200 rpm (OD
600
1.4).
The SWB media recipe [33] contains 342 mM NaCl, 14.8 mM MgCl
2
:6H
2
O, 0.1
mMCaCl
2
:2H
2
0, 6.7 mMKCl, 10 mMNH
4
Cl, 1 mMNa
2
SO4, 1 mM phosphate
buer (pH = 6.5) and was supplemented with 5 mM sodium acetate (electron
donor), 5 mM potassium nitrate (electron acceptor), vitamins, and trace metals
[151] to support anaerobic growth. Cells from this anaerobic pre-growth step
(which reached 8 x 10
8
CFU/ml) were then harvested by centrifugation at 7197
x g, washed 3 , concentrated in 3 ml of fresh SWB media without electron donor
or acceptor, and subsequently used to inoculate bioreactors for electrochemical
measurements.
3.2.2 Bioelectrochemical Measurements
The electrochemical characterization of Thioclava electrotropha ElOx9 was per-
formed in standard three-electrode bioreactors. A PW06 carbon cloth (Zoltek, St.
Louis, MO) measuring 2.5 cm x 1.5 cm was used as a working electrode, Pt wire
(CH Instruments, Austin, TX, USA) as a counter electrode, and Ag=AgCl (1M
KCl) (CH Instruments, Austin, TX, USA) as a reference electrode. The reactor
contained 50 ml of SWB media, supplemented with 25 mM sodium bicarbonate
(carbon source) and 5 mM nitrate (electron acceptor). Anaerobic conditions were
maintained by constantly purging the reactor with ltered and humidied 80:20
(v/v) N
2
=CO
2
gas mixture. Chronoamperometry and cyclic voltammetry (CV)
were performed using an 8 channel potentiostat (CH Instruments, Austin, TX,
USA). For chronoamperometry experiments, the working electrode was poised at
-278 mV vs SHE to act as electron donor for the cells. For CV, the working
28
potential was cycled between -378 mV to 622 mV vs SHE at a scan rate of 1
mV/s. Current was normalized to the projected surface area of the carbon cloth
electrodes. The reactors were inoculated with 4% (v/v) of the resuspended cell
culture once a stable baseline current was achieved; the cell density in the reactor
immediately after inoculation was 6.2 x 10
7
CFU/ml. To determine if CO
2
was
indispensable for electrochemical experiments, bicarbonate was removed from the
media and anoxic conditions in the bioreactor were maintained by purging with
pure N
2
.
Analysis of the electrode-attached biolms was performed after the cathodic
current in the bioreactor maximized and a CV was recorded. For media exchange
experiments, the bioreactor was opened in an anaerobic chamber to avoid interfer-
ence fromO
2
. The electrode (with attached biolm) was removed from the reactor
and swirled gently in SWB base media to remove loosely attached cells and wash
out old media caught in the carbon cloth electrode. It was then transferred to
a sterile bioreactor with fresh SWB media and CV was performed following this
exchange.
To perform electrochemical measurements of spent media, planktonic cells were
removed from reactor cultures either by ltering through 0.2m lter or by pellet-
ing and repeatedly centrifuging the supernatant in order to retain potential medi-
ators (7197 x g, 5 minutes, 10 times as described in [183] yielding spent media).
Spent media was then injected at 4% (v/v) in a bioreactor with pristine carbon
cloth poised at -278 mV vs SHE to test for soluble redox active shuttles. CV
was performed on the electrode two hours after being held under potentiostatic
conditions.
29
To perform electrochemical measurements under non-turnover conditions (no
electron acceptor), the bioreactor and carbon cloth electrodes were treated iden-
tically as the media exchange experiments described above, except that the elec-
trode/biolm was placed in fresh reactor SWB media without nitrate. The reactor
was maintained at -278 mV vs SHE under these non-turnover conditions for four
days, with CVs performed every 24 hours. In the experiments conducted to detect
any role for hydrogenases, after a CV on electrode attached biolm was performed
in 80:20 (v/v) N
2
=CO
2
, the atmosphere of the bioreactor was switched to 80:20
(v/v) H
2
=CO
2
. The electrode was poised at -278 mV vs SHE while being purged
with the H
2
rich gas for 1 hour prior to performing another CV.
3.2.3 Microscopy
Electrode-attached biolms were visualized using Scanning Electron Microscopy
(SEM) and Fluorescence Microscopy. For SEM, each carbon cloth electrode was
xed using 2.5% glutaraldehyde and serially dehydrated, in ascending concentra-
tions of ethanol (10%, 25%, 50%, 70%, 95% and 100%). The electrode was then
subjected to critical point drying and sputter coating prior to imaging at 15 keV
using JEOL JSM 7001F eld-emission microscope. For
uorescence microscopy,
bers of carbon cloth electrode were xed with 2.5% glutaraldehyde and incubated
in 5 g/ml of FM 4-64FX membrane stain (Molecular Probes, Life Technologies,
Inc) for 20 minutes. The samples were imaged on a Nikon Eclipse Ti-E inverted
microscope using a TRITC excitation/emission channel (Nikon lter set G-2E/C).
3.2.4 Protein Quantication
Total protein attached to electrodes was quantied as described previously [81].
Carbon cloth samples were subjected to alkaline lysis and digestion in 200mM
30
NaOH at 100
o
C for 90 mins, vortexing every 15 minutes. The extracted proteins
were then measured with a Pierce BCA Protein Assay Kit (Thermo Scientic,
Carlsbad, CA, USA).
3.2.5 Nitrate quantication
Nitrate consumption in the bioreactors was quantied using ion chromatogra-
phy (Metrohm 850 Professional Ion Chromatograph (Metrohm, Riverview, FL)
equipped with Metrosep A SUPP 5, 250 4 mm column and conductivity detector
with 3.2 mM sodium bicarbonate, 1 mM sodium carbonate running buer contain-
ing 2.5 % acetonitrile (
ow rate 0.7 mL/min)). Samples were collected immediately
after inoculation and at the end of chronoamperometry, ltered through 0.2 m
lter, and stored at 20
o
C until analysis.
3.3 Results and Discussion
Onset and continual increase of cathodic current was immediately observed upon
inoculation of T. electrotropha ElOx9 into bioreactors containing working elec-
trodes poised at -278 mV vs SHE and nitrate as the sole electron acceptor (Figure
3.1). This current response was consistently observed in multiple experiments
(n=8), reaching a maximal -2.45 1.30 A=cm
2
within 30-48 hours compared
to abiotic (sterile electrode) controls (at -0.05 0.05 A=cm
2
), indicating that it
results from cellular activity coupling cathodic electron uptake to nitrate reduc-
tion. The applied potential at the working electrode (-278 mV vs SHE) was chosen
to be well above that necessary for H
2
evolution in seawater [30, 151], to avoid
H
2
serving as a primary electron donor to the cells. To allow the electrode to
serve as electron donor to the cells, the chosen potential is also 100 mV lower than
31
Figure 3.1: Cathodic activity of Thioclava electrotropha ElOx9. Chronoamperom-
etry measurements show an increase in cathodic current after inoculation of cells
into a bioreactor containing a working electrode poised at -278 mV vs SHE. No
activity is detected from an uninoculated abiotic control (black).
potentials relevant to biological oxidation of zero valent sulfur in sea water, one of
the natural mineral electron donors for T. electrotropha ElOx9.
To further conrm that the current response resulted from metabolic activity,
the bioreactors were treated with 5 mM sodium azide, a known inhibitor of nitrate
reductase [126]. Introduction of azide resulted in a drastic, and subsequently sus-
tained, drop in cathodic current to -0.16 0.02 A=cm
2
(n=3) over the course
of 20 hours (Figure 3.2A). The observed inhibition, down to the level of abiotic
controls (Figure 3.2B), conrms the requirement of a functional nitrate respira-
tory pathway to aid the
ow of electrons from cathode. When not treated with
exogenously inhibitor, the observed maximal current decreased gradually over the
course of 14 days. Previous characterization of T. electrotropha ElOx9 revealed
it reduces nitrate to nitrite, but that its genome lacks a complete denitrication
pathway (GenBank accessionGCA
0
02085925:1) [30]. Since accumulation of nitrite
32
Figure 3.2: Eects of inhibition on cathodic activity. a) Chronoamperometry of
Thioclava electrotropha ElOx9, with a working electrode poised at -278 mV vs SHE,
shows cathodic activity and subsequent collapse of this activity after inhibition
with 5 mM azide. b) Summary (n=8) of cathodic currents detected from abiotic
(sterile) electrodes, maximal current after addition of T. electrotropha ElOx9 cells,
and as a result of inhibition with 5 mM azide.
to high concentrations is known to inhibit cellular activity [145], it is possible that
the accumulation of this respiratory product under our batch reactor conditions is
responsible for the gradual decrease in electron uptake observed over two weeks.
While our initial experiments contained bicarbonate and CO
2
, to allow for
potential autotrophic growth, we also tested whether this was necessary for the
observed cathodic activity of T. electrotropha ElOx9. Omission of this inorganic
carbon source in the media still resulted in the usual current response upon inoc-
ulation (Figure 3.3). Though it is generally dicult to exclude all carbon sources,
this observation suggests the increasing cathodic activity over time is primarily
associated with increasing cellular attachment to the electrodes and/or adaptation
to the electrode as an energy source, rather than increasing overall biomass from
cellular growth and division. In addition to direct visualization with microscopy
(see below), cellular attachment was conrmed by measuring the total protein
33
content attached to the electrode at the end of the chronoamperometry measure-
ments and found to be 0.10 0.03 mg=cm
2
(n=3). Following a previous example
[81], the total protein content and observed cathodic current were used to heuris-
tically estimate the average per cell EET rate into T. electrotropha ElOx9. Using
a cellular volume of 0.36 m
3
(from Scanning Electron Microscopy) and represen-
tative protein content of 0.2 g/ml [121], the inward EET rate amounted to 1.4
x 10
4
e

=s=cell, a gure roughly one to two orders of magnitude lower than out-
ward EET rates measured in the anode-respiring microbe S. oneidensis MR-1 [64].
Cellular nitrate reduction resulted in a 0.3 mM decrease in nitrate concentration
during the chronoamperometry measurements. Comparing the total Coulombs
transferred from the cathode (by integrating the current over time) to the maxi-
mum possible Coulombs, based on 2 electron reduction of all the consumed nitrate
to nitrite, reveals a Coulombic eciency of 43.9%. Biocathodes dominated by
Thiobacillus sp., reducing nitrate to nitrite, have been previously reported to yield
comparable coulombic eciencies of 24.5% - 46.9% [140].
To shed some light on the mechanism(s) of electron uptake by T. electrotropha
ElOx9 from the electrode, cyclic voltammetry (CV) was performed once maximal
current was reached in chronoamperometry measurements. Under turnover condi-
tions, (i.e. in the presence of nitrate as an electron acceptor) a reversible catalytic
wave was observed with an onset potential of 236 mV (Figure 3.4). This catalytic
current, which was absent in abiotic (sterile media) controls, re
ects sustained
electron transfer from the electrode to nitrate through cells and continuous regen-
eration of the proteins in this cellular electron transport chain [113]. By examining
the rst derivative of the turnover CV (Figure 3.5), the location and width at half
max (110 8 mV) of the peak associated with the catalytic wave were found to
34
Figure 3.3: Cathodic activity of Thioclava electrotropha ElOx9 without carbon
source. Chronoamperometry of T. electrotropha ElOx9 cells in the absence of any
inorganic carbon source (no bicarbonate in SWB reactor media, and no CO
2
in
purging gas). On inoculation of cells in the bioreactor, with carbon cloth poised at
-278 mV vs SHE, cathodic current increases to similar levels as seen in experiments
with inorganic carbon sources.
be consistent in multiple experiments, despite variation in overall current magni-
tude. Since an ideal redox-active thin layer is expected to exhibit 90/n mV peak
width [155], and allowing for the expected broadening that results from biolms
containing both inert components and heterogeneous redox proteins with dispersed
potentials and rate constants [183], it is likely that the electrochemical reaction
detected involves a single electron (n=1) transfer process.
To assess whether extracellular electron uptake by T. electrotropha ElOx9
involves a direct contact mechanism or mediation via soluble shuttles, turnover
voltammetry was also performed on electrodes/biolms after exchange with fresh
media lacking planktonic cells or previously secreted factors. Following media
exchange, the sigmoidal catalytic waveform was largely retained with only a small
loss in cathodic current when compared to measurements prior to the media
35
Figure 3.4: Turnover Cyclic Voltammetry (CV) of Thioclava electrotropha ElOx9.
Turnover CV (in presence of nitrate) of T. electrotropha ElOx9 reveals a catalytic
wave indicating sustained extracellular electron uptake from electrodes coupled to
nitrate reduction by the cells. The catalytic wave is retained after exchanging old
media with fresh media (purple). CVs of cell-free spent media shows no detectable
redox features.
exchange (Figure 3.4). The small dierence is likely attributed to some cell detach-
ment from the electrodes during electrode rinsing prior to placement in fresh media.
To further investigate any possible contribution from diusible redox molecules to
cathodic activity, cells were removed from the spent media of bioreactors and this
spent media was subsequently injected in new bioreactors with sterile working
electrodes. Voltammetry performed with this spent media showed no detectable
diusion-controlled redox features (Figure 3.4).
To visualize cellular attachment and the extent of biolm formation on elec-
trodes, the working electrodes were imaged using scanning electron microscopy
(SEM) and
uorescence microscopy using the membrane stain FM 4-64FX (Fig-
ure 3.6). Both imaging techniques revealed a dispersed monolayer of attached
cells, with occasional dense clusters. Cells were observed as rod-shaped and 1-2
36
Figure 3.5: First derivative of turnover cyclic voltammetry (CV). a) Representative
derivative CV from both forward and reverse scans. The width at half-max is 105
mV. b) Derivative CVs of multiple experiments (with varying cathodic activity)
showing similar location and width at half-max of 110 8 mV. Only reductive
scan derivatives are shown in (b) for clarity.
m in length, consistent with the expected morphology of T. electrotropha ElOx9
[30]. Taken collectively, cellular attachment to the electrodes, retention of cathodic
activity following fresh media exchange, and the absence of redox activity from sol-
uble components point to a contact-based, rather than diusible shuttling, extra-
cellular electron uptake process.
Non-turnover conditions (i.e. in the absence of nitrate as an electron acceptor)
were also used to assess the redox activity of T. electrotropha ElOx9 components
on electrodes. Voltammetry under these non-turnover conditions allows an inves-
tigation of interfacial electron transfer in a manner less concealed by high catalytic
currents and independent of additional confounding factors such as diusive limi-
tations of species into cells or the respiration rate of the microbes [114]. Following
measurements under turnover conditions (described above), the working electrodes
were transferred to reactors lacking nitrate and again maintained at an applied
37
Figure 3.6: Microscopic characterization of electrodes. a) Scanning electron
microscopy of carbon cloth electrode bers after electrochemical measurements of
Thioclava electrotropha ElOx9 cells (Scale bar 10 m). Inset shows higher magni-
cation of the same electrode and attached cells (Scale bar 2m). b) Fluorescence
microscopy image of the carbon cloth electrode bers shows attached cells using
FM 4-64FX to stain cell membranes (Scale bar 10 m).
potential of -278 mV vs SHE. After four days the cathodic current decayed to
baseline level and CVs showed a transition from the catalytic wave to reversible
peaks (Figure 3.7), revealing the redox center with a formal midpoint potential at
-94 mV. Furthermore, to conrm that the nitrate deprivation step did not alter the
physiology of T. electrotropha ElOx9, nitrate was reintroduced to the reactor after
non-turnover measurements. Re-introduction of nitrate restored catalytic activity
to 91% of that observed prior to removal (Figure ??), indicating no permanent
harm to the cells from the four-day nitrate deprivation step.
A recently demonstrated cathodic electron uptake mechanism (by
methanogens) involves the extracellular release of cell-produced hydroge-
nases that attach to electrodes and catalyze H
2
evolution, which in turn functions
as electron carrier to cells [39, 153]. Given the ability of T. electrotropha ElOx9
38
Figure 3.7: Comparing turnover and non-turnover voltammetry. Turnover cyclic
voltammetry (CV) of electrode-attached Thioclava electrotropha ElOx9 cells (red)
compared to an abiotic control (black). Removal of nitrate results in a non-turnover
CV (purple) revealing redox peaks indicated by arrows. Catalytic activity is re-
gained by the cells upon re-introduction of nitrate (blue).
to grow on H
2
as an electron donor [30], we set out to assess whether such as a
hydrogenase catalyzed mechanism might play a role in our study. Voltammetry
of electrode attached hydrogenases is known to exhibit reversible behavior (both
proton reduction and hydrogen oxidation) [7] with the expected formal potential
of the 2H
+
=H
2
couple at -384 mV vs SHE at pH 6.5. The midpoint of the
catalytic wave observed in this study is considerably higher, suggesting that
hydrogenases are not involved in facilitating electron uptake. To conrm this
hypothesis, we performed an H
2
addition experiment, as previously described
[153]. Introduction of H
2
can help dierentiate between H
2
-dependent (by
detecting current associated withH
2
oxidation) andH
2
-independent mechanisms.
No H
2
oxidation current was detected after purging with H
2
for an hour (Figure
3.8), further ruling out involvement of hydrogenase in facilitating electron uptake
39
Figure 3.8: Eect of hydrogen addition. Turnover cyclic voltammetry of electrode-
attached Thioclava electrotropha ElOx9 cells before (red) and after (blue) purging
with 80% H
2
=CO
2
for 1 hour (blue).
by T. electrotropha ElOx9 under our experimental conditions. A small decrease
in overall catalytic activity was however observed, relative to before H
2
addition,
which is consistent with the introduced H
2
acting as a competitive electron donor
to cells instead of the cathode serving as sole electron source.
3.4 Conclusion
This electrochemical investigation demonstrates that the recently isolated sulfur-
oxidizing marine bacterium Thioclava electrotropha ElOx9 is capable of extracel-
lular electron uptake from solid-state electrodes and coupling this functionality to
nitrate respiration under anaerobic conditions. By studying the electrochemical
activity of the cells (amperometry and voltammetry under both turnover and non-
turnover conditions), eects of media exchange, and analysis of spent media, our
40
measurements indicate that T. electrotropha ElOx9 performs this inward extra-
cellular electron transfer through a direct contact-based mechanism via a redox
center with a formal potential of -94 mV vs SHE, rather than soluble intermedi-
ate electron carriers. While the molecular identity of this extracellular electron
conduit is not yet known, the recently available draft genome of T. electrotropha
ElOx9 points to some intriguing candidates, including transmembrane mono- and
multi-heme c-type cytochromes that may function in an analogous manner to the
known bidirectional electron conduits in metal-reducing bacteria [71, 152]. The
electrochemical activity demonstrated here motivates future studies to investi-
gate dierentially expressed genes under cathodic and S
0
oxidizing conditions,
targeting selected genes for mutations, and further electrochemical characteriza-
tion of mutants to denitively pinpoint the identity of the charge carriers. Given
its cathodic activity, metabolic
exibility, and emerging genomic information, T.
electrotropha ElOx9 is well positioned to serve as a promising model system for
understanding inward extracellular electron transfer.
41
Chapter 4
Extracellular Electron Transfer in
the Human Microbiome
4.1 Introduction
Extracellular electron transfer (EET) is a metabolic strategy used by microorgan-
isms to harness energy by interfacial electron
ow to or from abiotic surfaces (e.g.
minerals or poised electrodes) [62, 159]. EET is accompanied by generation of a
proton motive force (PMF) comprising of a change in pH and membrane potential
across the cellular membrane thus driving ATP synthesis [122]. Microbes like She-
wanella oneidensis MR-1 , isolated from anaerobic sediments of lake Oneida, New
York [127] and Geobacter sulfurreducens, isolated from the freshwater sediments of
Potomac river, Maryland [107] were the rst microbes reported to be capable of
outward EET, allowing them to respire solid mineral electron acceptors. Detailed
characterization of these microbes has led to our current understanding of out-
ward EET mechanisms. Specialized protein conduits spanning across the bacterial
membrane facilitate outward electron transfer by direct contact [24, 49, 147] or cell
secreted redox molecules (like
avins) act as electron shuttles mediating the elec-
tron transfer [112, 171]. This phenomenon, however, is more extensive as a wide
range of electroactive organisms spanning both bacterial and arcehael domains have
been detected in a variety of environments [23, 55, 77, 78, 81, 148, 151, 157, 167].
42
Electron transfer and energy conservation is applicable to all of biology includ-
ing microbes beyond natural or engineered environments. Some of the preliminary
evidences of EET activity in human associated microbes was observed in microbial
fuel cells treating wastewater and conductive laments in biolms causing necrotic
infection of jaw bones [77, 78, 176]. We hypothesize that diverse microorganisms
associated with the anaerobic niches of the human body [43] may be capable of
performing EET by interacting with insoluble particulate matter or epithelial cells
to meet their energy demands.
We rst focus on a model lung bacterium, Pseudomonas aeruginosa PA14, to
investigate EET mechanism in the mammalian lung environment. P. aeruginosa
is an opportunistic pathogen that is known to cause chronic infections in patients
suering from cystic brosis (CF) by forming thick biolms with persister cells
that are resistant to antibiotics [4, 101, 177]. These microbes also produce colorful
redox active molecules called phenazines [141] that act as a virulence factor [100]
providing immunity against competing microorganisms [13], serving as a quorum
sensing signal [41], controlling colony morphology and biolm formation [40], and
promoting iron acquisition [175]. Importantly, phenazine redox cycling has been
shown to promote anaerobic survival [59] and redox hemostasis [142] in P. aerug-
inosa. In the presence of an external electron acceptor, phenazines can mediate
EET by shuttling electrons between the cells and the electron sink, hence enabling
anaerobic survival [174]. This process may be relevant in the oxygen deprived areas
of CF lung biolms where higher concentrations of phenazines have been detected
[179, 182].
While the details of EET shuttling mechanism by phenazines are still unclear,
a recently published study has implicated the cytoplasmic
avo-proteins of pyru-
vate and-ketoglutarate dehydrogenase in the reduction of phenazine [61] enabling
43
ATP synthesis during glycolysis . Reduction of phenazines require 2e

and 2H
+
at neutral pH, implying that release of reduced phenazine into extracellular space
is likely accompanied by the generation of PMF [60] causing hyperpolarization of
the cellular membrane. Eux pumps like MexAB-OprM and MexEF-OprN have
been reported to be involved in the release of these shuttles [181]. The reduced
phenazines transfer electrons to an external electron acceptor (e.g. anode) and
get oxidized before diusing back to the cells [174]. The extracellular process of
phenazine redox activity, especially on electrodes, has been more extensively stud-
ied. Many accounts exploiting phenazine synthesis and redox activity have found
applications in the eld of biotechnology, specically in the area of electrochemical
sensing and bioelectrochemical technologies like microbial fuel cells [136]. Reports
explaining the eects of cell strain, carbon source, quorum sensing cascade and
electrode potential on phenazine production and oxidation currents have also been
published [21, 22, 168].
The most relevant work focused at understanding the mechanism of extracellu-
lar electron shuttling by phenazines in P. aeruginosa using an external electrode
was published by Wang et al. [174]. This study undertook an electrochemical
approach using an electrode poised at an oxidizing potential which served as an
electron acceptor for cell-secreted reduced phenazines to enable shuttling. While
this study was important in establishing the role of both endogenous phenazines
and an external electron acceptor in promoting long term anaerobic survival in P.
aeruginosa PA14, it lacks sucient electrochemical data supporting the observa-
tions.
The human gut ecosystem, unlike the lung environment (represented by model
bacterium P. aeruginosa), is far less understood, mainly because of the lack of
candidate micoorganisms that can represent it. Many electrode enrichment studies
44
have been previously reported that led to successful isolation of bacteria represen-
tative of the environment being analyzed [81, 103, 151, 154]. A recently published
study applied a similar approach to fecal samples with an aim to isolate microbes
representative of human gut. Electrochemical enrichment at 0.4 V vs SHE fol-
lowed by incubation with lactate or acetate as electron donor and MnO
2
as the
electron acceptor led to the isolation of two electroactive bacteria { Enterococ-
cus avium and Klebsiella pneumoniae [129]. Separate studies are emerging that
are aimed at a detailed characterization of other isolates associated with human
gut, like, Enterococcus faecalis [89, 134], Listeria monocytogenes [102] and Faecal-
ibacterium prausnitzii [91, 92]. F. praunitzii is an anaerobe that has been shown
to utilize a
avin mediated shuttling pathway during EET. In case of E. faecalis,
reduced demethylmenaquinone (DMK) in the cytoplasmic membrane was found to
be essential for electron transfer to conductive surfaces, whereas heme and heme
proteins were not. L. monocytogenes also requires reduced membrane associated
DMK to transfer electrons to an external electron acceptor through a series of

avin based proteins. Understanding EET mechanism in these microbes is cru-
cial to not only support the currently lacking literature associated with EET in
Gram-positive bacteria but also, to reveal dierent metabolic pathways adopted
by microbes in the gut.
It is also imperative to recognize that most of the studies that account for our
current understanding of the mechanisms of electron transfer between cells and
electrodes stem from bulk electrochemical analyses. Bulk techniques overlook the
nuances and dynamics of sub-populations or even behavior of single cells that may
be contributing towards their electroacitvity. Recently, a technique was developed
45
in our lab to in vivo monitor EET activity in S. oneidensis MR-1 at a single-
cell level in a bioelectrochemical system (BES) by hyphenating it with
uores-
cence microscopy (Pirbadian et al., unpublished). A
uorescent dye, Thio
avin T
(ThT) was used to report membrane potential which was observed to be positively
correlated with electrode potential indicating that cells experience a bioenergetic
advantage when on poised electrodes.
In this chapter, with regards to P. aeruginosa, we aim to provide additional
details on phenazine electron shuttling mechanism by bridging the gaps from the
previous report [174], especially with respect to limited electrochemical data. We
use tools like chronoamperometry and cyclic voltammetry to answer other ques-
tions such as the eect of alternate electron acceptor on electron shuttling and the
in
uence of mixing in BES on the eciency of phenazine cycling. Additionally,
we also attempt to spatially and temporally resolve the bulk BES to understand
the dynamics of EET at a single-cell level by combining electrochemistry with
live-cell
uorescence imaging to monitor the behavior of membrane potential of P.
aeruginosa PA14 cells in response to applied potential.
On the gut microbiome front, in collaboration with the lab of Prof. Steven
Finkel, we take an exploratory approach to target electroactive microbes from a
community derived from human fecal samples using electrochemical enrichment by
using oxidized solid-state electrodes. This has led to the isolation of dierent bac-
teria, all exhibiting varying levels of electroactivity, hence, presenting themselves
as suitable candidates capable of serving as model organisms to study EET mech-
anisms in the human gut ecosystem. Eorts to isolate and electrochemically char-
acterize new microorganisms will contribute towards assessing the existing EET
models or to unravel new pathways enabling a deeper comprehension of metabolic
routes adopted by microbes in the gut environment.
46
4.2 Materials and Methods
4.2.1 Cell Growth Conditions
Pseudomonas aeruginosa PA14
For bulk electrochemical analysis, Pseudomonas aeruginosa PA14 (WT and phz)
was grown aerobically in 50 ml lysogeny broth (LB) from a frozen stock (80
o
C,
30% glycerol stock) at 37
o
C and 200 rpm for 18 hours till stationary phase was
attained (OD
600
1.9-2.1). The cells were harvested by centrifuging the entire cul-
ture at 7197 g for 10 minutes and washed 3 for 5 minutes and concentrated
in 1 ml of fresh AB media [34] containing 3 mM (NH
4
)
2
SO
4
, 8.4 mMNa
2
HPO
4
,
4.4 mMKH
2
PO
4
, 10.2 mM NaCl, 0.1 mMCaCl
2
:2H
2
O and 1 mMMgCl
2
:6H
2
O.
The resuspended culture was used to inoculate bioreactors for electrochemical mea-
surements.
For membrane potential measurements, cells were grown in 5 ml LB from a
frozen stock (80
o
C, 30% glycerol stock) at 37
o
C and 200 rpm for 24 hours. The
cells were harvested by pelleting and washing the culture at 7197 g, 3 for 5
minutes and resuspended in the same volume of fresh AB media.
Fecal community
Fecal samples for analysis were obtained from a nonprot stool bank, OpenBiome
(Cambridge, MA). The samples were collected from screened and proled stool
donors, processed to remove large particulate matter and delivered in 12.5% ster-
ile glycerol and 0.9% buered saline solution. Microbial community associated with
the stool sample (stored at 80
o
C) was outgrown in YCFA media modied from
[25]. The base media contains 2 g casitone, 1 g yeast extract, 52 mM NaHCO
3
,
47
2.5 mM K
2
HPO
4
, 3.3 mM KH
2
PO
4
and 15.3 mM NaCl with nal pH adjusted
to 7.4. Sterilized base media was supplemented with 0.05 mM MgSO
4
:7H
2
0, 0.05
mMCaCl
2
:2H
2
O, 0.0025 mM hemin, Wolfe's vitamin solution (from 1000 stock
solution), 0.5 mM glucose, 0.29 mM maltose, 0.29 mM cellobiose, 30 mM acetic
acid, 9 mM propionic acid, 1 mM butyric acid, 1 mM valeric acid and 1 mM iso-
valeric acid. Sample was inoculated in 20 ml YCFA media in test tubes, then
purged with N
2
for 20 minutes and sealed using a rubber stopper. Microaerobic
conditions were maintained by piercing the stopper with a 21G needle before incu-
bation at 37
o
C on a roller for 48 hours. The needle also disabled accumulation of
toxic by-products in the tube head-space. The cells were harvested by pelleting
and washing the culture at 7197 g, 3 for 5 minutes and resuspended in 1 ml
of fresh YCFA base media. This culture was used to inoculate bioreactors for elec-
trochemical measurements. Isolates were also cultured and harvested using same
conditions. Determination of cell density (via spread plating) and isolation of sin-
gle colonies by repeated streaking was performed on YCFA agar plates prepared
from YCFA media with 1.5% agar. Isolation of microbes from biolm community
was accomplished by gently sonicating the electrode-attached biolm in 2 ml of
YCFA base media and spreading the community on an agar plate. Distinct and
individual colonies from these plates were picked and streaked on fresh plates until
an isolated colony was obtained. All isolates were saved in 10% glycerol in YCFA
base media and stored at 80
o
C.
4.2.2 Bioelectrochemical Measurements
The electrochemical characterization was performed in standard three-electrode
bioreactors. A PW06 carbon cloth (Zoltek, St. Louis, MO) measuring 1.0 cm x
1.0 cm was used as a working electrode, Pt wire (CH Instruments, Austin, TX,
48
USA) as a counter electrode, and Ag/AgCl (1M KCl) (CH Instruments, Austin,
TX, USA) as a reference electrode. Chronoamperometry and cyclic voltammetry
(CV) were performed using an 8 channel potentiostat (CH Instruments, Austin,
TX, USA). Current was normalized to the projected surface area of the carbon
cloth electrodes.
Pseudomonas aeruginosa PA14
The reactor contained 50 ml of AB media, supplemented with 20 mM glucose as the
electron donor and 20 M pyocyanin. Anaerobic conditions were maintained by
constantly purging the reactor with ltered and humidiedN
2
gas. For chronoam-
perometry experiments, the working electrode was poised at 622 mV vs SHE to
act as electron acceptor for the cells. For CV, the working potential was cycled
between -378 mV to 722 mV vs SHE at a scan rate of 1 mV/s. The reactors were
inoculated with 1% (v/v) of the resuspended cell culture once a stable baseline
current was achieved; the cell density in the reactor immediately after inocula-
tion was 3.5 10
9
CFU/ml. Small aliquots (0.1 ml) were withdrawn from the
reactor at intervals of approximately 24 hours to determine cell density over the
course of 7 days using serial dilution. The eect of nitrate addition was studied by
injecting 25 mM potassium nitrate in the bioreactor 5 hours after cell inoculation.
For obtaining cell free inoculum, planktonic cells were removed from LB culture
by pelleting and repeatedly centrifuging the supernatant to retain any released
redox active compounds (7197 g, 5 minutes, 15 times as described in [183]. This
inoculum was then injected at 1% (v/v) in a bioreactor with carbon cloth poised
at 622 mV vs SHE. Experiments without gas purging or stirring were performed in
an anaerobic chamber by assembling the reactors aerobically and purging with N
2
49
for 1.5 hours before transferring in the chamber. The inoculum was also purged
for 15 minutes before injection to avoid oxygen interference.
Fecal community
The reactor contained 50 ml of YCFA media lacking casitone, yeast extract, hemin
and ribo
avin from vitamins. Anaerobic conditions were maintained by constantly
purging the reactor with lteredN
2
gas. For chronoamperometry experiments, the
working electrode was poised at 422 mV vs SHE acting as an electron acceptor.
The reactors were inoculated with 1% (v/v) of the resuspended cell culture once
a stable baseline current was achieved; the cell density in the reactor immediately
after inoculation was 10
8
CFU/ml. For lysed cell control, the reactor inoculum
was sonicated at high amplitude for 1-2 minutes, occasionally cooling it in an ice
bath. For heat-killed control, the cells were placed in a hot water bath (60
o
C) for
30 minutes before injecting in the bioreactor.
4.2.3 Membrane Potential Analysis
Bioreactors were constructed using a cylindrical glass tube glued to a transparent
and patterned gold coated electrode (coating thickness 5 nm Ti and 10 nm Au
on glass). To enable electrical contact with the electrode, a piece of copper wire
was attached using silver paint (protected by epoxy glue). Customized reactor
top contains slots to hold Pt counter electrode, an Ag/AgCl reference electrode
and a port for gas inlet, outlet and sampling. Total capacity of these bioreactors
is around 10-12 ml. Resuspended cells were transferred to a sterile reactor and
allowed to remain unperturbed for 10 minutes, enabling some of them to settle on
the electrode. The planktonic culture was then replaced by 8 ml sterile AB media
containing 20 mM glucose, 20 M pyocyanin and 10 M Thio
avin T. This was
50
done in order to reduce background
uorescence from planktonic cells. The reactor
was placed on a Nikon Eclipse Ti-E inverted microscope and purged constantly
with humidied N
2
to maintain anaerobic conditions. Chronoamperometry was
performed by poising the gold electrode at 622 mV vs SHE using a Wavedriver 20
Bipotentiostat/ Galvaostat, Pine Research, NC. Green
uorescence of cells over 18
hours was determined by time-lapse microscopy using a FITC excitation/emission
channel (Nikon lter set B-2E/C) with 500 ms exposure time. Image processing
for quantication of
uorescence intensity and plotting was done using MATLAB
(MathWorks).
4.3 Results and Discussion
4.3.1 Electrochemical investigation of electron shuttling
mechanism in Pseudomonas aeruginosa PA14
An increase in anodic current was observed upon inoculation of P. aeruginosa
PA14 wild-type (WT) in a bioreactor containing a working electrode poised at 622
mV vs SHE and glucose as electron donor (Figure 4.1A). Over 72 hours, the current
sustained at 0.42 0.09A=cm
2
(n=3), over the abiotic (sterile electrode) control
(at 0.21 0.02 A=cm
2
) and did not increase over time. While P. aeruginosa
PA14 is capable of synthesizing phenazine carboxylic acid (PCA) under anaerobic
conditions [146], the steady current indicates that under the experimental condi-
tions of bioreactor operation, cells did not synthesize PCA over the observed length
of time or that the concentration of PCA produced by the cells is not enough to
be detected by the poised electrode. Phenazines are colored compounds whose
51
Figure 4.1: Electrochemical analysis of Pseudomonas aeruginosa PA14 wild-type
(WT). a) Chronoamperometry measurements indicate an increase in anodic cur-
rent upon inoculation of P. aeruginosa PA14 WT cells in a bioreactor containing
a working electrode poised at 622 mV vs SHE. The current remains steady over
50 hours indicating no detectable amount of phenazines building up over time. b)
Cyclic voltammetry (CV) after 50 hours reveal no signicant change in electro-
chemical activity when compared to abiotic control (black).
presence can be visually observed by detecting color change in P. aeruginosa cul-
tures. The color of bioreactor culture remained unchanged throughout the course
of the experiment, also indicating the absence of phenazines. This observation
is re-armed by cyclic voltammetry (CV) which shows no signicant dierence
between PA14 (WT) and abiotic control (Figure 4.1B). Hence, to acquire a better
control over the concentration of phenazine in the bioreactor, I decided to work
with a strain lacking the phenazine biosynthesis genes, phz, with a known amount
of pyocyanin (a derivative of PCA [141]) exogenously added.
Chronoamperometry of P. aeruginosa PA14phz with added pyocyanin shows
a steady increase in anodic current upon inoculation in a bioreactor with a working
electrode poised at 622 mV vs SHE and glucose as the electron donor (Figure
4.2A). The anodic current density consistently reached a maximum of 18.5 4.4
52
A=cm
2
(n=6) over 24 hours before gradually decreasing over 90 hours. The color
of phenazines is also indicative of their redox state[60]. The reactors are initially
blue with oxidized pyocyanin during cell inoculation but turn colorless as more
reduced pyocyanin accumulates over time, possibly due to low rate of phenazine
re-oxidation at the electrode. Sterile control with only pyocyanin and control with
only phz cells, showed no such anodic activity implying that both pyocyanin and
cells contribute to an increase in current. Since phz cells did not interact with
the working electrode, the increase in anodic current density can be attributed
to the oxidation of pyocyanin at the electrode after undergoing reduction by the
cells. The passivation of the electrode by formation of an inert biolm possibly
causes a gradual decline in the current over time. CV of pyocyanin reveals a
reversible wave feature centered at -21 mV under the experimental conditions,
which is absent in the control without pyocyanin. The diminishing current density
observed in chronoamperometry at the end of 90 hours with cells in the bioreactor
is re
ected in the CV as well. Another reversible peak with mid-point potential of
-165 mV is also observed whose origin is currently not known (Figure 4.2B).
To test the metabolic activity of cells during electron transfer to the electrode,
glucose was excluded from the bioreactors. Following inoculation of phz cells,
anodic current increased and reached a maximal density of 19 3.74A=cm
2
which
is comparable to the current observed in bioreactors with glucose (Figure 4.3A).
To rule out rich media components potentially being carried over from overnight
culture and acting as electron donor, cells were washed ve times instead of three
before inoculation, however, the current density remained unaected. Polyhydrox-
yalkanoates (PHA) are polymers produced by Pseudomonas as reserved carbon
and energy source under stressed growth conditions [84] and could be serving as
53
Figure 4.2: Electrochemical analysis of Pseudomonas aeruginosa PA14phz. a)
Chronoamperometry measurements indicate an increase in anodic current upon
inoculation of P. aeruginosa PA14phz cells in a bioreactor containing a working
electrode poised at 622 mV vs SHE with 20M pyocyanin (blue). No activity was
detected in an uninoculated abiotic control containing only pyocyanin (black) or
inoculated control containing only cells (purple). b) Cyclic voltammetry (CV) of
pyocyanin indicates a reversible peak with mid-point potential at -21 mV (black).
No activity is observed without pyocyanin (purple). Suppressed current density
with cells and pyocyanin (blue) after 90 hours possibly occurs due to electrode
passivation.
a carbon source indicating that glucose may not be the only carbon and electron
source for the cells in the bioreactor.
To conrm whether cells used the working electrode as a terminal electron
acceptor (TEA), nitrate was added in the bioreactor. Nitrate serves as an alter-
nate electron acceptor for P. aeruginosa under anaerobic conditions [19, 142].
Addition of nitrate in the bioreactor led to a rapid decline in the anodic current
indicating a switch in cellular metabolism from anode reduction to nitrate reduc-
tion. The preference of cells to use nitrate as the TEA could be explained by its
high formal potential for 2e

reduction (430 mV) compared to pyocyanin (-21 mV)
(Figure 4.3B). The color of planktonic reactor media remained blue throughout the
experiment indicating that pyocyanin was not reduced by the cells.
54
Figure 4.3: Eect of glucose and nitrate on the anodic activity of Pseudomonas
aeruginosa PA14phz. a) Comparable anodic current densities detected from
cells in bioreactors with 20 mM electron donor glucose (blue) and without glucose
(purple). b) Addition of 25 mM nitrate in order to substitute for the poised
working electrode as an electron acceptor for the cells, led to a rapid decline in
anodic current (red) as compared to the bioreactor without any nitrate (blue).
Figure 4.4: Anodic activity of cell-free inoculum of Pseudomonas aeruginosa
PA14phz. Chronoamperometry of cell-free inoculum of P. aeruginosa PA14phz
indicates no increase in anodic current upon inoculation in a bioreactor contain-
ing an electrode poised at 622 mV vs SHE (red) compared to the abiotic control
(black).
55
Figure 4.5: Analysis of planktonic cell density. Summary (n=3) of bioreactor
planktonic cell density of P. aeruginosa PA14phz, expressed in colony forming
units per ml (CFU/ml), compared between Day 1 (day of bioreactor inoculation,
dark gray) and Day 7 (light gray), under dierent conditions { with cells, 20 M
pyocyanin, 20 mM glucose and working electrode poised at 622 mV vs SHE),
without poised working electrode, without added pyocyanin (PYO) and without
added glucose. A slight loss in planktonic cell density is observed in all conditions
but the rst.
I next proceeded to conrm that the increase in anodic current was not a result
of lysed cell products released in the inoculum during washing. Cells were removed
from the inoculum by repeated centrifugation before injecting in the bioreactor.
The resulting current density remained unchanged and reached 2.16 0.53A=cm
2
after 45 hours, comparable to the sterile abiotic control (Figure 4.4).
The planktonic cell density in bioreactors was compared between day 1 (the day
of inoculation) and day 7 to investigate whether the cells experienced any long-
term metabolic advantage when provided with an oxidized electrode, pyocyanin
and glucose. When provided with all three parameters, the cell density moderately
56
increased by 1.67 10
8
CFU/ml from the initial cell density of 7.9 x 10
9
CFU/ml.
On the other hand, when either one of the factors { poised electrode, pyocyanin
or glucose were not provided to the cells, their density declined by 1.33 10
8
CFU/ml, 5 10
8
CFU/ml and 1.20 10
9
CFU/ml, respectively (Figure 4.5). This
decline in cell density, while expected, was not as drastic as reported previously
[174]. This could be attributed to the dierent media composition used for our
experiments, especially the concentration of ammonium (15 mM instead of 93 mM)
[90]. It is possible that depriving the cells for a duration longer that 7 days may
lead to a greater drift between the initial and nal cell densities, however, under
the present experimental conditions of the bioreactor, other mechanisms enabling
cellular sustenance are likely at play.
Anodic current density was observed to be positively correlated with the mixing
of bioreactor planktonic culture (Figure 4.6A). Under static condition (no nitro-
gen purging or stirring) in an anaerobic chamber, the maximum current density
obtained was 3.18 0.7 A=cm
2
. However, actively purging the bioreactor with
humidied nitrogen led to a higher current density of 18.5 4.4 A=cm
2
. Fur-
ther, introducing stirring to a purging bioreactor resulted in an even higher anodic
current density at 43.2 4.4 A=cm
2
. This indicates that ecient mass transfer
of reduced pyocyanin to the working electrode leads to higher anodic current den-
sity [11]. Subsequently, we used these current densities to calculate the number of
times pyocyanin could cycle between the cells and the poised electrode. One com-
plete redox cycle would be dened as the cellular reduction of oxidized pyocyanin,
followed by its re-oxidation at the electrode and diusion back to the cells to get
reduced (inset, Figure 4.6B). The frequency of these cycles for a 2e

redox reaction
of pyocyanin (20M concentration in a 50ml bioreactor volume) over 77 hours was
determined by integrating the area under chronoamperometry plots. The average
57
Figure 4.6: Eects of stirring on the eciency of pyocyanin cycling. a) Chronoam-
perometry of P. aeruginosa PA14phz shows a dependence of anodic current den-
sity on the extent of mixing in the bioreactor. The current density is lowest in
static bioreactors (red), higher in actively purged bioreactors (blue) and highest in
actively purged and stirred bioreactors. b) The frequency of pyocyanin redox cycles
between P. aeruginosa PA14phz cells and the working electrode increased with
enhanced mixing of planktonic culture in the bioreactor. At 77 hours of chronoam-
perometry, the number of cycles calculated for a static bioreactor increased from
an average (n=3) of 5 cycles to 13 (with N
2
purging) and to 29 (with purging and
stirring).
number of cycles calculated for a static bioreactor increased from 5 cycles to 13
(with N
2
purging) and to 29 (with purging and stirring). Following these gures,
and approximate planktonic cell density of 10
9
CFU/ml, electron transfer rates
per cell were heuristically calculated for all three conditions. EET rates accounted
for 2.2 10
4
e

/cell/s for static bioreactor, 6.4 10
4
e

/cell/s for bioreactor with
N
2
purging and 1.3 10
5
e

/cell/s for bioreactor with N
2
purging and stirring.
Additionally, while not experimentally tested, EET rates will also rely on the con-
centration of pyocyanin. These calculated respiration rates are approximately one
to two orders of magnitude lower than outward EET rates measured in S. onei-
densis MR-1 [64]. These results imply that the extent of phenazine redox cycling
58
Figure 4.7: Experimental set-up for the analysis of cellular membrane potential.
a) Schematic representation of the bioreactor- cylindrical glass body with optically
transparent patterned gold coverslip at the base as a working electrode, reactor top
with counter and reference electrode. Cells settled on the working electrode are
stained from Thio
avin T
uorescent dye in reactor media and observed through
an inverted microscope. b) Patterned gold electrode used for the experiment -
glass coverslip coated with 5nm titanium and 10 nm gold.
relies on the perturbation in the environment of study. It has been shown that
cystic brosis (CF) patients have restricted blood and gas exchange in their lungs
due to plugged airways from excessive mucus [98], likely keeping the system barely
perturbed. Considering this in the context of our experiments, CF lungs would
loosely resemble static cultures, hence the metabolic advantage to the cells due to
phenazine redox shuttling may be low.
To enable live-cell imaging of P. aeruginosa PA14 cells on the electrode with
simultaneous electrochemical measurements, a three-electrode bioreactor with an
optically transparent and patterned gold electrode was placed on an inverted epi
u-
orescence microscope (Figure4.7). The patterned electrode allowed us to monitor
cell
uorescence on both the electrode and glass, in parallel. To measure whether
the cells can generate an active proton motive force (PMF), a
uorescent cationic
dye Thio
avin T (ThT) was used to report membrane potential. This dye has been
59
successfully shown to report membrane potential in Bacillus subtilis [143] and S.
oneidensis MR-1 (Pirbadian et al., unpublished). I conrmed that this dye can
also reliably respond to the membrane potential in P. aeruginosa cells. This was
established by bubbling oxygen in a reactor along with 10 M ThT and observing
an increase in
uorescence intensity over time corresponding to an increase in the
PMF. The
uorescence intensity immediately collapsed upon addition of 100 M
carbonyl cyanide m-chlorophenyl hydrazone or CCCP (a protonophore [87]) by
leveling proton gradient across the cellular membrane (Figure 4.8C). Under anaer-
obic conditions, with the working electrode poised at 622 mV for 18 hours and with
20 M pyocyanin exogenously added, the
uorescence response of ThT indicated
that the PMF did not increase over time. This may imply a very low membrane
potential gain that doesn't increase over time but sustains to enable cell survival
(Figure 4.8D). When the
uorescence intensity on poised electrode was compared
to the intensity on glass in the same experiment, it was observed that the cells on
the electrode barely outdid those on glass in terms of membrane potential (Fig-
ure 4.8E). This observation could be explained by the low electron transfer rates
(calculated above) generating a low PMF, that likely makes it dicult for ThT to
respond to. Electron transfer rates, calculated for carbon cloth used in the bulk
electrochemical experiments, is already low, and may be even lower for planar gold
electrodes used in these experiments [85] further aecting the PMF. PAO1 strain
uses the eux pump MexGHI-OmpD for the extracellular release of pyocyanin [2]
but absence of this pump in PA14 strain did not aect its discharge [156]. This
implies that reduced pyocyanin may not require an eux pump and should be able
produce a PMF, or, it may use other eux pumps, that, if require active trans-
port (require ATP), will not be able to generate enough PMF. While these claims
have not been experimentally veried, depending upon which condition holds true
60
Figure 4.8: Eect of applied potential on the membrane potential of P. aeruginosa
PA14phz. a) Cells under aerobic conditions display high Thio
avin T (ThT)

uorescence intensity before addition of 100 M carbonyl cyanide m-chlorophenyl
hydrazine (CCCP) (scale bar 5 m). b) Cells under aerobic conditions display
reduced Thio
avin T (ThT)
uorescence intensity after addition of 100M CCCP
(scale bar 5 m). c) ThT
uorescence intensity of cells increase over time under
aerobic condition followed by a sudden decrease after the addition of CCCP. d) No
increase in ThT
uorescence (green) or current (orange) was observed over time
with a constant applied potential of 622 mV vs SHE (purple). e) Comparable ThT

uorescence of cells on the poised electrode (green) and on glass (black) indicates
no signicant gain in membrane potential for cells from the applied potential.
for pyocyanin release, PMF and hence, the cellular membrane potential will be
aected and will in
uencing ThT
uorescence.
Through extensive electrochemical studies, we conrm that pyocyanin can act
as a redox shuttle that enables EET in P. aeruginosa PA14. We were also able to
quantify the extent of phenazine shuttling and deduce that it depends upon the
environment of the cells (perturbed vs unperturbed). Whether cells experience
61
Figure 4.9: Electrochemical activity of metacommunity human fecal sample.
Chronomaperometry measurements indicate an increase in anodic current upon
inoculation of the fecal community in a bioreactor with a working electrode poised
at 422 mV vs SHE (red). Addition of 10 mM cyanide, a respiratory chain inhibitor,
led to a rapid decrease in the anodic current (gray). No activity was observed in
the sterile abiotic control (black).
any energetic gain from a poised electrode is less clear probably because of low
rates of electron transfer.
4.3.2 Electrochemical enrichment and isolation of gut
microbes from human fecal samples
Fecal samples for the enrichment of gut microbes were obtained from a non-
prot stool bank, OpenBiome (Cambridge, MA). The samples were collected from
healthy individuals, processed to remove particulate matter and pre-screened for
infectious viruses, parasites, bacteria and antibiotic resistant bacteria by the stool
bank. We outgrew these samples microaerobically, in a complex rich media [25]
designed to mimic the human gut environment. Facultative microbes cultivated
in this media were introduced into a bioreactor with a working electrode poised at
422 mV vs SHE leading to an increase in anodic current over 5 hours to a maximal
62
Figure 4.10: Anodic activity of lysed and heat killed samples. a) Chronoamperom-
etry of fecal community samples either lysed (blue) or heat-killed (purple) before
inoculation in a bioreactor with working electrode poised at 422 mV vs SHE pro-
duced no activity compared to regular inoculum. Lysed community control showed
an in increase in current after 45 hours. b) Chronoamperometry of all three colonies
isolated from lysed community control led to an increase in anodic current upon
inoculation in bioreactors poised at 422 mV vs SHE (red, blue, purple). No activity
was detected in the sterile abiotic control (black).
current density of 4.73 1.0 A=cm
2
, followed by a gradual decrease over next
45 hours (Figure 4.9). Increase in current density was observed against a sterile
abiotic control (0.04 0.01 A=cm
2
). Addition of 10 mM cyanide, a common
respiratory chain inhibitor [67] led a rapid decline in the anodic current density to
0.15 0.04A=cm
2
, comparable to the abiotic control. This implies that the rise
in current is biological and can be attributed to cell respiration and/or activity of
redox proteins.
Additional proof supporting the presence of a cell-anode interaction was
observed by introducing heat-killed cells in the bioreactor which did not lead to
an increase in anodic current density compared to the untreated inoculum (Figure
4.10A). To determine whether the electrochemical activity is derived from redox
63
molecules secreted by lysed cells, all cells in the inoculum were lysed by sonica-
tion and injected in the bioreactor. Resulting current density remained close to
the abiotic control implying lysed cell products were not responsible for the biore-
actor activity. The lysed cell control bioreactor maintained low currents for 30
hours before increasing and attaining a maximum of 0.8 A=cm
2
. This increase
in current density was accompanied by an increase in cell density (10
3
CFU/ml
to 10
5
CFU/ml). Three newly emerged and morphologically distinct colonies were
isolated (all three isolates were identied as dierent strains of E. coli through
16s rRNA sequencing) and subjected to further electrochemical characterization.
Chronoamperometry showed that the anodic current density for the isolates peaked
at 0.71, 0.96 and 1.22A=cm
2
in 17.5, 18 and 35 hours, respectively, demonstrating
electroactivity of the isolates (Figure 4.10B). It is interesting to note that E. coli
is capable of exporting
avins (a knowne

shuttle for S. oneidensis) with the help
of a multidrug and toxic compound extrusion protein, YeeO [117], however, the
uptake of
avins by cells is not possible [178]. Only an exhaustive study to deter-
mine EET mechanisms in these strains will help us conclude whether shuttling or
other EET routes can provide any metabolic gain to the cells.
Bioreactor with metacommunity fecal sample led to the formation of a polymi-
crobial biolm on the electrode. This was observed by plating lightly sonicated
electrode-attached biolm in buered media on agar displaying seven distinct
colonies that diered from each other based on size, opacity and morphology.
Based on anodic current density observed in community sample bioreactor, it is
reasonable to expect some of these microbes were enriched on the electrode due to
some selective respiratory advantage oered by the poised electrode. Based of 16S
rRNA sequencing three of these colonies were identied as E. faecalis, E. coli and
64
Figure 4.11: Anodic activity of isolated colonies. Chronoamperometry of three
colonies, Enterococcus faecalis (red), Escherichia coli (blue) and Klebsiella pneu-
moniae (purple), isolated from electrode attached biolm of fecal community indi-
cated an increase in anodic current upon inoculation in a bioreactor containing a
working electrode poised at 422 mV vs SHE.
K. pneumoniae. On inoculating these isolates in a bioreactor with a working elec-
trode poised at 422 mV vs SHE, an increase in anodic current density was observed.
Approximately 8 hours after inoculation, peak current density of 0.98, 1.07 and
1.64A=cm
2
was observed for E. faecalis, E. coli and K. pneumoniae, respectively
(Figure 4.11). Other colonies isolated from the biolm are yet to be identied.
While a detailed characterization of electron transfer mechanisms by these isolates
is underway, the preliminary chronoamperometry results show promise in a suc-
cessful exploitation of electrode enrichment in detecting and isolating electroactive
candidate microorganisms representing the human gut microbiome.
Enterococcus faecalis is a fermentative lactic acid bacteria commonly found
in the mammalian intestine [58]. The microbe does not have cytochromes as it
cannot synthesize hemes. However, when provided with extracellular hemes, it
can assemble cytochromes and undergo respiration by glucose oxidation and oxygen
65
reduction via dimethylquinone [14, 134, 180]. E. faecalis is gaining interest as a
model organism for detailed EET characterization because it is a Gram-positive
microbe. Unlike Gram-negative bacteria, EET in these microbes is less understood
since they lack an outer membrane and possess a thick (20-35 nm) [170] non-
conductive peptidoglycan layer. From studies conducted on E. faecalis OG1RF, it
was concluded that the microbe can undergo direct electron transfer with graphite
electrodes or indirectly via osmium redox polymers. It was also inferred that
cytochromes, which are essential for EET in many microbes [32], actually impede
outward electron transfer in E. faecalis [134]. Another study on strain ZER6,
highlighted that E. faecalis can undergo EET via redox shuttling using ribo
avin
[186]. This is interesting, since a study on E. faecalis (another Gram-positive
bacteria) recently uncovered the presence of a new
avin based EET mechanism
where electrons from dimethylmenaquinone can be picked up by a series of
avin
based proteins and exported to the outside of cell surface [102]. Such mechanism
could also be present in E. faecalis and further investigation is required. E. faecalis
may also be able to undergo syntrophic interactions within the gut community with
microbes that produce or require
avins (e.g. E.coli or F. prausnitzii)[91, 117].
4.4 Conclusion
Electrochemical investigation of P. aeruginosa PA14 (phz) reveals that for this
bacterium, electron transfer with an external anode is facilitated by the cycling
of exogenously added small redox molecules called phenazines. Quantication of
total charge transferred to the electrode with the help of a xed concentration of
phenazines indicated that the eciency of redox cycling directly depends upon the
mixing in the bioreactor system. Lowest number of redox cycles were recorded
66
for static bioreactors with an increase observed for purged reactors, followed by
purged and stirred reactors. Membrane potential response of bacteria on poised
electrodes to detect any energetic gain experienced by the cells is still unclear due
to low
uorescence signal, likely due to low rates of electron transfer. Anodic
enrichment of fecal bacteria from human fecal samples gave rise to a current signal
that was armed to be biological in nature. Subsequently, isolation and identi-
cation of microbes from electrode attached biolms revealed three electroactive
microbes representing the human gut environment. A detailed investigation of
electron transfer pathways of these microbes can have important implications in
understanding the unexplored metabolic pathways adopted by them.
67
Chapter 5
Bacterial Immobilization for
Atomic Force Microscopy under
Physiological Conditions
5.1 Introduction
Atomic force microscopy (AFM) is a scanning probe technique that generates
images based on short range interactions between a sample surface and a posi-
tion sensitive probe (sharp tip mounted on a cantilever). The resolution of AFM
images depends upon the radius of the probe or tip [12]. Current success in tip
fabrication allows for sub-nanometer resolution which makes AFM ideal for study-
ing both structural and molecular properties of microorganisms [45]. Single-cell
AFM analysis has not only helped visualize cell structure dynamics and deduce its
morphology [46, 139, 165], but also to assess and quantify properties, such as, adhe-
sivity and elasticity [1, 166], and identify certain chemical groups and molecular
recognition sites [35, 57].
AFM techniques have also assisted in advancing our understanding of extracel-
lular electron transfer (EET). EET is a metabolic strategy adopted by microbes
that enables them to respire external solid electron acceptors (e.g. minerals or
anodes) by expelling electrons across an insulating cell membrane with the help of
redox proteins called outer membrane cytochromes [159]. One of the mechanisms
68
of EET involves electron conduction across long membrane laments or pili [72].
Charge transport properties across these structures have been analyzed in She-
wanella oneidensis MR-1 [49], Geobacter sulferreducens [99], Geobacter uraniire-
ducens[164] and Rhodopseudomonas palustris RP2 [169] using conductive { AFM,
in which a bias is applied between the tip and sample to measure current across the
sample. Antibody recognition force microscopy revealed that S. oneidensis MR-1
outer membrane cytochromes, OmcA and MtrC, are expressed on the cell surface
when they are grown on iron (III) oxide as a terminal electron acceptor [109]. In a
separate study, AFM aided in concluding the formation of specic bonds between
hematite and S. oneidensis MR-1 cytochromes [108].
In contrast to high resolution electron microscopy where samples must be ana-
lyzed under vacuum following harsh sample preparation, AFM provides the user

exibility to control the environment of the sample. Samples can be analyzed in
air under dry conditions or in liquid under physiological conditions. Imaging under
air is a popular choice since sample preparation is easy and cells adsorb on the
solid surfaces (e.g. on glass slides or mica) which makes scanning easy [95]. Many
of the electron transport studies in EET microbes have been performed under dry
conditions. The advantage of doing these studies under liquid with live cells is that
they would be physiologically more relevant. Specialized tips with insulated can-
tilevers oering low leakage currents now being available, studying electron transfer
properties of microbes under liquid is more feasible [52]. Techniques like AFM {
scanning electrochemical microscopy (AFM { SECM) can now be performed, that
can undertake a correlative topographical and electrochemical investigation of sin-
gle cells under physiological conditions [51]. In case of SECM, the amperometric
tips give rise to a current response based on the substrate topography and reac-
tivity with a resolution depending upon the size of the tip. Conventional SECM
69
enables scanning of the substrate at a constant height. Hence, when both substrate
reactivity and topography are variable, it is dicult to resolve the two convoluted
components [110]. Alternatively, AFM allows mapping the topography of the sub-
strate by interactions between cantilever tip and the sample giving a nanometer
scale resolution. It only provides topographical image but does not allow detection
of any functionalization (like enzymes, photoactive or redox active molecules) on
the biological sample. The main advantage of integrating AFM and SECM is that
the AFM component of the tip can be used for high-resolution (nanometer scale)
topographical imaging or for maintaining a desired tip/substrate separation, and
the SECM component can electrochemical characterization of biological samples
in liquids [5].
However, one of the major challenges of performing live-cell imaging using
AFM lies in the ecient immobilization of cells on a substrate. Cells are prone
to detachment in liquid because of their small size that provides only a small sur-
face contact area and due to lateral forces exerted by tip during scanning [47].
Live cell motility and low adhesion also poses an issue towards keeping the cells
stable during imaging [95]. As a result, a number of strategies have been devel-
oped and implemented to immobilize cells including physically trapping them in
membrane lters or microwells, modifying surfaces using specic coatings to enable
localization of cells either by electrostatic interactions or via formation of covalent
bonds. The eectiveness, advantages and disadvantages of these immobilization
techniques has been previously reviewed [118]. It should be noted that there is
not one immobilization strategy that works for all cells because various structural
(shape or size) and chemical properties of the cell dictates ecacy of the technique
70
[95, 118]. Regardless, a sturdy and reproducible method allowing long term sta-
bility of the cells during imagining while also keeping them viable is required to
explore techniques like AFM { SECM.
In this chapter, I explore dierent immobilization techniques to determine a
robust one that works consistently on the three phylogenetically diverse electroac-
tive microbes examined in this dissertation { Shewanella oneidensis MR-1, Pseu-
domonas aeruginosa PA14 and Thioclava electrotropha ElOx9. I determined that
surface modication using polycationic amino acid (poly L-lysine) led to a stable
connement of cells on the surface via glutaraldehyde cross-linking. We further
assessed the eects of this procedure on cell structure, size and viability, conrming
that the cells were not harmed as a result of the immobilization. Finally, I eval-
uated the extent of adhesion of the three microbes by comparing phase contrast
images. In tapping mode AFM, amplitude of an oscillating tip changes from the
free air amplitude depending upon the interactive forces. The change in amplitude
is monitored to yield a topographical map of the surface. In addition to the ampli-
tude, phase of the tip comparative to the driver response also changes when the
tip experiences an area of dierent material compositions as shown in (Figure 5.1)
[115]. This shift in phase dierence gives rise to contrast images. The contrast
depicts various material properties of the sample under investigation, including
adhesion [119].
5.2 Materials and Methods
5.2.1 Cell Growth Conditions
Shewanella oneidensis MR-1 and Pseudomonas aeruginosa PA14 were grown aer-
obically in 5 ml Lysogeny Broth (LB) from a frozen stock (80
o
C, 30% glycerol
71
Figure 5.1: Illustration of phase response of an AFM tapping tip on a sample with
dierent material properties. As the tip is scanned across a sample, the phase of
the oscillating tip (red) relative to the driver signal (black) changes when on the
surface compared to when on the cells.
stock). S. oneidensis MR-1 was incubated at 30
o
C and P. aeruginosa PA14 at
37
o
C and 200 rpm for 24 hours. Thioclava electrotropha ElOx9 was grown aero-
bically in 5 ml LB + ions (LBI) media which contains (per liter) 10 g tryptone,
5 g yeast extract, 20 g NaCl, 3 g MgCl
2
:6H
2
O and 0.15 g CaCl
2
:2H
2
O, from a
frozen stock (80
o
C, 30%) at 30
o
C and 200 rpm for 24 hours. The cells grown in
rich media were harvested by centrifugation at 7197 g and washed 3 in phos-
phate buered saline (PBS) which contains (per liter) 8 g NaCl, 0.2 g KCl, 1.44 g
Na
2
HPO
4
, 0.24 g KH
2
PO
4
with pH adjusted to 7.2. Cells were resuspended in
the same volume (5 ml) of PBS before depositing it on the substrate. For ElOx9,
modied PBS with 20 g/L NaCl was used.
5.2.2 Substrate Preparation
Silicon dioxide wafers were used for analysis as a substrate. Approximately 0.5
0.5 cm size wafer was cleaned by sonication in approximately 5 ml of acetone,
isopropyl alcohol, 70% ethanol and deionized (DI) water for 10 minutes each.
The cleaned wafers were dried under a stream of ltered N
2
and stored until use.
72
Cleaned wafer was placed in a 12-well plate and approximately 0.5 ml of Poly-
L-lysine (PLL) was added to the well or enough to cover the wafer. It was then
allowed to soak in PLL for at least one hour at room temperature or left overnight
at 4
o
C. PLL was removed from the well by aspiration using a pipet tip. The wafer
was rinsed twice with 1ml DI water. 1 ml of 5% glutaraldehyde was added in the
same well and the wafer was left to soak overnight. Glutaraldehyde was removed
by aspiration and the wafer was washed 10 with 1 ml DI water.
5.2.3 Sample Preparation
The wafer was transferred to a clean well in the 12-well plate. 1 ml of the washed
and resuspended cell culture was placed on the prepared substrate and cells were
allowed to settle and attach for 15 minutes. The wafer was gently rinsed 3 with
300 l PBS to wash o weakly bound cells without letting it dry before imaging.
To ensure attachment, another sample prepared in parallel was allowed to dry
completely under a stream of ltered N
2
.
5.2.4 Atomic Force Microscopy
Sample imaging was done using a Cypher ES Environmental AFM (Oxford Instru-
ments, Asylum Research, Santa Barbara, CA). Images were obtained by tap-
ping mode (AC water mode). Small silicon nitride cantilever with a silicon tip,
(BioLever Mini, BL-ACT04S, Oxford Instruments, Santa Barbara, CA) with a res-
onant frequency f = 110 Hz in air (around 35 Hz in liquid) and stiness k = 0.09
N/m, was used for this study. The cantilever was driven to its resonant frequency
using a piezo. All images were taken under PBS or modied PBS using a liquid
perfusion cantilever holder with a completely sealed environment, at 30
o
C. A scan
rate of 0.75 to 1 Hz was used for imaging.
73
5.2.5 Evaluation of cell viability
Cell viability in terms of membrane integrity was assessed using BacLight
LIVE/DEAD staining kit (Invitrogen), containing two nucleic acid stains SYTO9
and Propidium Iodide (PI). Both stains were added to 1 ml of washed and resus-
pended cell culture to a nal concentration of 1.67M SYTO9 and 2M PI. This
culture was then placed on the prepared substrate and left unperturbed for 15
minutes in the dark, allowing the cells to stain while getting settled and immo-
bilized. The sample was then washed 3 with 300 l PBS and placed upside
down on a clean glass coverslip. The samples were imaged on a Nikon Eclipse Ti-E
inverted microscope using a TRITC excitation/emission channel (Nikon lter set
G-2E/C) and FITC excitation/emission channel (Nikon lter set B-2E/C). Images
were merged and processed using ImageJ.
5.2.6 AFM Image Processing
Image processing on AFM images was performed using Gwyddion software. Flat-
tening lter was applied on the images and the substrate tilt eect was removed
by using polynomial t lter. The height and phase of the silica substrate was set
to 0 before quantication using line proles.
5.3 Results and Discussion
Several methods of bacterial immobilization were tested to image live Shewanella
oneidensis MR-1, Thioclava electrotropha ElOx9 and Pseudomonas aeruginosa
PA14 cells using AFM under liquid. To support cells, a biocompatible and inert
surface is required that does not undergo microbial degradation [124]. Commonly
74
used and cheap inorganic substrates such as glass coverslips or silica wafers were
considered. Given the ease of focusing the sample on an opaque surface under
a liquid droplet, we proceeded with using silicon wafer as a substrate instead of
transparent glass slides.
To check whether surface modication for cell immobilization could be avoided,
biolms were allowed to form naturally on the silica wafer by placing it in a plank-
tonic culture. Though cell attachment was observed, their distribution over the
surface was uneven and imaging was dicult, probably due to cell stacking which is
expected in the early phases of biolm development [94]. One of the common and
more reproducible techniques involving no surface treatment of the substrate, but
only physical entrapment of cells in membrane lters was also ruled out because
it works best for coccoid bacteria of size comparable to the lter pores [47], and
all the microbes under consideration for this study are rod-shaped Gram-negative
bacteria. Hence, other procedures more inclusive of cell shape and size, were ana-
lyzed.
Coating substrate with poly-L-lysine (PLL) and gelatin have been shown to
localize cells on the surface successfully [42, 104]. PLL is a polycationic polymer of
lysine that strongly adsorbs on a surface, exposing its cationic groups [116]. Cell
surfaces of Gram-negative microbes, that are negatively charged due to lipopolysac-
charides on their outer membrane [111], immobilize on PLL coated surfaces due
to electrostatic interactions. Gelatin is also composed of a mixture of dierent
amino acids and relies on electrostatic and hydrophobic interactions to immobilize
cells [42, 118]. Samples that were allowed to dry out after immobilization and
gentle washing step, showed cloudy deposit on the surface indicating cells adhered
to the surface [3]. However, wet samples that were imaged using tapping mode
AFM, showed several loosely bound cells that dislodged during imaging. High
75
ionic strength of media aects the electrostatic forces that hold the cells in place.
It is possible that ionic strength of phosphate saline buer (PBS) weakened the
interaction between the coated slides and cells, aecting their immobilization [104].
5.3.1 Surface modication and analysis
Another tool to localize cells on a surface is via formation of covalent bonds.
Glutaraldehyde, a bifunctional aldehyde, is commonly used as a crosslinker between
the surface and many biomolecules [124]. The schematic representation of the
immobilization procedure used in our study is illustrated (Figure 5.2). We used
PLL coated wafers with exposed primary amines. The amines are protonated
under pH 10.2 and can undergo Schi's reaction with glutaraldehyde forming an
imine (C=N) [135]. The other aldehyde group undergoes the same condensation
reaction with amines on the cell membrane serving as a link between cells and
the amine-functionalized surface [124]. Some of the studies that have used this
crosslinking technique, have proceeded via activation of all amines on the cell
surface by introducing glutaraldehyde to the entire cell culture [65, 86]. This will
aect the surface chemical properties of the entire cell and viability of the culture.
The eects of glutaraldehyde xation are minimized in this procedure, where the
surface is activated rst, and cells are deposited on the modied surface after.
Hence, crosslinking on cells occur only at the area of contact, thereby preserving
the overall properties of the cell.
The eect of chemical modication on silicon surface was observed by imaging
a small surface area (500 nm) of the wafer as described previously [65]. The
amplitude images reveal a dierence in roughness between the uncoated (Figure 5.3
A) and the coated (Figure 5.3 B) silicon surface. This observation was conrmed
by collecting a height prole from height images of the wafer across the frame
76
Figure 5.2: Schematic representation of immobilization technique. a) Silicon wafer
coated with Poly-L-lysine (PLL) with exposed amines. b) Condensation reaction
between exposed amines and glutaraldehyde leads to the formation of imine bond.
c) Amines on cell surface of microbes undergo another condensation reaction with
glutaraldehyde leading to crosslinks and cellular immobilization.
(Figure 5.3 C, D) displaying larger variations in the coated surface. A common
parameter used to determine the roughness of a surface area is the root mean
square roughness or Sq which is dened as the root mean squared deviation from
the mean. Sq for the two surfaces was determined using the image processing
software Gwyddion and calculated to be 1.146 nm for uncoated surface and 1.718
nm with PLL/glutaraldehyde modication conrming surface modication.
5.3.2 Assessment of structure, size and viability of immo-
bilized cells
Cells for AFM analysis were obtained from early stationary phase cultures in rich
media. Three phylogenetically diverse and electroactive microbes, S. oneidensis
MR-1, T. electrotropha ElOx9 and P. aeruginosa PA14 were used for this study.
The cells were imaged both in air and in liquid (PBS buer). Figure 5.4 shows
77
Figure 5.3: AFM analysis of silicon substrate. a) AFM image of uncoated silicon
wafer (Scale bar 100 nm). b) AFM image of silicon wafer coated with Poly-L-lysine
(PLL) and glutaraldehyde (Scale bar 100 nm). c) Height prole of uncoated silicon
wafer. d) Height prole of silicon wafer coated with PLL and glutaraldehyde.
amplitude images of these microbes prepared and imaged under the two conditions.
One of the striking observations on comparing the cells imaged in air vs in liquid
is the dierence in the surface morphology. Dehydrated cells have a wrinkled
appearance. On the other hand, morphology of cells immobilized and imaged
under liquid is smooth and featureless. Air drying can cause deformation in the cell
membrane due to loss of water resulting in wrinkling, cracks,
attening and other
artifacts [42]. Hence, ultrastructures that are visible on dried cell surfaces, are not
visible under liquid. P. aeruginosa PA14 imaged under air (Figure 5.4 C) reveal

agella that are not visible under liquid. This could be due poor immobilization of
the ne structures causing their movement by lateral forces exerted by the tip [42].
Size analysis on cells was performed using line proles on height images (Figure
78
Figure 5.4: Comparison of AFM images of cells under dry and under liquid con-
ditions. Tapping AFM amplitude images of a) Shewanella oneidensis MR-1 b)
Thioclava electrotropha ElOx9 c) Pseudomonas aeruginosa PA14 on silicon wafer
under dry conditions. (Scale bar 4 m). Tapping AFM amplitude images of
d) S. oneidensis MR-1 e) T. electrotropha ElOx9 F) P. aeruginosa PA14 cells
immobilized on silicon wafer and imaged under phosphate buered saline under
physiological conditions (Scale bar 1 m).
5.5). Length and diameter of MR-1 cells ranged from 2 - 7 m and 0.5 { 1.0 m;
of ElOx9 cells from 1.7 { 2.5 m and 0.7 - 1.0 m; of PA14 cells from 1.3 { 1.7
m and 0.9 { 1.1 m, respectively. These measurements agree with previously
reported values for MR-1 [33], ElOx9 [30] and PA14 [56].
Both PLL (polycationic amino acid) and glutaraldehyde (common xative) are
cytotoxic in nature and can aect the viability of immobilized cells [44, 73]. There-
fore, to check whether reagent toxicity and/or the formation of covalent bonds on
cell surface have harmed the cells, we applied LIVE/DEAD staining technique with
nucleic acid stains SYTO9 and Propidium Iodide (PI) as previously described [104].
The viability assessment was performed based on the cellular membrane integrity
79
Figure 5.5: Height analysis of cells under liquid conditions. AFM height images
and corresponding line proles of a) Shewanella oneidensis MR-1 b) Thioclava
electrotropha ElOx9 and c) Pseudomonas aeruginosa PA14 displaying height across
the drawn white line. (Scale bar 1 m).
80
Figure 5.6: Viability assessment of immobilized cells. LIVE/DEAD staining of a)
Shewanella oneidensis MR-1 b) Pseudomonas aeruginosa PA14 and c) Thioclava
electrotropha ElOx9 indicating membrane integrity of the cells (Scale bar 5 m).
and permeability. Cells with intact membrane are stained green by SYTO9 and
were considered living, whereas PI permeates the cells through compromised cell
membranes and stains them red, indicating dead cells. Fluorescence images of
immobilized and stained cells reveal that most of the cells were alive following the
localization procedure (Figure 5.6). This was ensured by thoroughly washing the
substrate with de-ionized water after modication before depositing cells on them
for immobilization, to get rid of any residual free PLL or glutaraldehyde.
Overall these results indicate that PLL/glutaraldehyde modication to sili-
con wafer enabled consistent immobilization of cells via covalent bonding enabling
AFM imaging under liquid droplet. The images revealed smooth cell surface struc-
ture unlike in dehydrated cells. Cells did not appear deformed and their sizes were
comparable to previously reported values. Finally, LIVE/DEAD staining indicated
that the chemical modications that led to cell localization, did not aect their
viability and hence, they are t for physiological studies.
81
5.3.3 Phase Image Analysis
In addition to providing topographical maps, tapping mode AFM can also pro-
duce phase contrast images. These images are generated due to the phase dier-
ence between the oscillation signal driving the piezo and the oscillation signal of
cantilever. While height and amplitude images are useful in analyzing the surface
structure of the cells, information about their physical properties, such as adhesion,
friction and elasticity can be deduced using phase images. Phase images can also
resolve any compositional heterogeneity (including static charge density) present on
the surface [119]. As per convention, a phase dierence of 90
o
was set between the
excitation and response oscillation signal when the cantilever was driven at the res-
onance frequency [115]. Closer to the surface, the change in oscillation amplitude
leads to the de
ection of phase from 90
o
. This de
ection of phase angle translates
into the energy dispersing onto the sample from the tip, which in turn provides a
measure for adhesion, friction, elasticity or electrostatic nature of the sample [115].
It should be noted that phase contrast can arise from any interaction between the
tip and the sample, all explaining dierent properties of the cell. To isolate the
cause of phase dierence, further experimentation is required that will enable us
to single out the type of tip-sample interaction causing the phase dierence. In
this study, we hypothesize that the phase contrast images explain adhesiveness of
the cells under investigation. Higher the magnitude of phase shift, higher is the
energy dissipated, implying that the sample is more adhesive. A discrete color
scheme is used to highlight the phase contrast in comparison to the substrate for
S. oneidensis MR-1 (Figure 5.7 A), T. electrotropha ElOx9 (Figure 5.7 B) and P.
aeruginosa PA14 (Figure 5.7 C), caused likely by cell adhesion. A contrast from a
white/blue substrate background towards pink/black cells represents a de
ection
82
Figure 5.7: Qualitative assessment of adhesion. Phase contrast images with dis-
crete color scheme representing cell adhesion of a) Shewanella oneidensis MR-1,
b) Thioclava electrotropha ElOx9, c) Pseudomonas aeruginosa PA14 (Scale bar 1
m)
in phase angle caused by the presence of cells on the surface, indicating the extent
of energy dispersion and hence, the extent of adhesion. While this is only a quali-
tative representation, it was observed that cells with lower phase contrast (yellow
or green) tended more to get dislodged from the surface during the raster scan
(represented by arrows in Figure 5.7). However, cells with higher contrast (pink)
could be imaged for at least 1.5 hours without any movement suggesting better
adhesion. It can therefore be suggested that the extent of cell adhesion is one
of the contributing factors towards the phase dierence, indicating the success of
PLL/glutaraldehyde immobilization technique. It is estimated that the surface of
a single S. oneidensis MR-1 cell is covered with 10
3
10
4
e

transfer proteins called
outer membrane cytochromes. Since interaction of the tip with charged surfaces
can also contribute towards phase shift, it will be interesting to tease apart elec-
trostatic contribution towards phase contrast by testing the cytochrome decient
mutant of MR-1 cells and comparing it with wild-type.
Quantitative measurements of phase contrast were accomplished by calculating
the dierence between phase angle at the substrate and at the cell surface. These
83
Figure 5.8: Extent of cell adhesion. AFM phase images and corresponding line
proles of a) Shewanella onedensis MR-1 b) Thioclava electrotropha ElOx9 and c)
Pseudomonas aeruginosa PA14 displaying shift in phase across the drawn white
line (Scale bar 1 m). d) Mean phase shifts over several cells observed in S.
oneidensis MR-1, T. electrotropha ElOx9 and P. aeruginosa PA14.
84
values were obtained by drawing line proles across several cells as shown in (Figure
5.8). Negative phase shift is observed due to more attractive interaction of the tip
on the soft cell compared to the hard silicon substrate [115]. The magnitude of
average phase shift was calculated to be 16.14 1.33 degrees for S. oneidensis MR-
1, 20.96 1.6 degrees for T. electrotropha ElOx9 and 9.80 1.84 degrees for P.
aeruginosa PA14. Assuming cell adhesion to be major factor responsible for phase
contrast, one can infer from this observation that the immobilization technique
worked slightly variably for the three Gram-negative microbes. Lower adhesion in
P. aeruginosa PA14 can be explained by high production of extracellular polymeric
substance (EPS). It is counter-intuitive since EPS is known to facilitate biolm
formation, however if cells are deposited on the modied silicon wafer, the EPS will
settle on the surface before the cells, thereby forming a `conditioning lm' [36]. This
can block exposed aldehyde groups on the silicon wafer. Surfaces pre-coated with
EPS have also been shown to negatively aect cell attachment [75]. Additionally, it
was tedious to wash the overnight culture before deposition, because, unlike ElOx9
and MR-1, PA14 cells did not form a tight pellet and may have led to carry-over
of some EPS. Higher phase contrast observed in ElOx9 cells could be due to their
smaller size. Lateral forces experienced by larger MR-1 cells during raster scanning
by the tip could aect adhesion.
5.4 Conclusion
In the present study, I showed that phylogenetically diverse microbes focused
in this thesis { S. oneidensis MR-1, T. electrotropha ElOx9 and P. aeruginosa
PA14 were successfully immobilized on amine functionalized silicon wafers using
glutaraldehyde without aecting the cell viability. Additionally, phase contrast
85
imaging enabled a comparison between the extent of adhesion of each of the cell
types. Immobilization of a diverse range of bacteria has important implications
in exploiting their characteristics by AFM under physiologically relevant condi-
tions. Bacterial localization also constitutes the rst step towards exploring more
complicated techniques like AFM-SECM.
86
Chapter 6
Conclusion
A diverse group of microorganisms exhibit a unique capability of using insoluble
minerals or electrodes as electron donors or acceptors for their metabolic gain.
This process is known as extracellular electron transfer (EET) and has impor-
tant implications in understanding various biogeochemical cycles and enhancing
microbial electrochemical technologies. While EET mechanisms are well under-
stood in dissimilatory metal reducers, the process is functionally diverse (micro-
bial metal oxidation) and ubiquitous (observed in natural and engineered envi-
ronments). This thesis contributes towards understanding the reverse process of
electron uptake from reduced electron donors. We also explore the extent the
EET process in host-associated microbes that have not been well-explored so far.
Finally, a bacterial immobilization technique was developed that would enable
atomic force microscopy (AFM) studies under liquid and development techniques
like AFM { scanning electrochemical microscopy (AFM-SECM) further allowing
characterization of single-cell electron transfer properties of electroactive bacteria.
In chapter 3, a detailed electrochemical characterization of Thioclava elec-
trotropha ElOx9 electron transfer mechanisms under anaerobic conditions success-
fully demonstrated that the cells rely on a direct-contact based electron uptake
mechanism facilitated by a redox moiety centered at -94 mV vs SHE. The multi-
heme cytochrome conduit of Shewanella oneidensis MR-1 responsible for anode
reduction has also been reported to be involved in cathode oxidation. Since
the T. electrotropha ElOx9 draft genome contains several mono and multiheme
87
cytochromes [30] similar to the ones found in S. oneidensis MR-1 [152], it will be
interesting to test the bidirectionality of the redox entities involved in the elec-
tron uptake by these cells. Another important task would be to identify the redox
proteins involved in the electron uptake pathway of T. electrotropha ElOx9 to
better understand their electron transfer properties. This can be accomplished
by dierentially expressing genes during cathode and sulfur oxidation and target-
ing specic genes for mutations, followed by identifying functions of the mutated
genes. Whether T. electrotropha ElOx9 cells experience any appreciable bioener-
getic gain on a poised cathode is also an important question and can be studied
using
uorescent membrane potential indicators such as Thio
avin T (ThT) [143].
The
uorescence response of the probe with respect to the applied potential will
reveal if cells can produce any proton motive force (PMF) on the electrode.
In chapter 4, electrochemical enrichment of microbes from human fecal sam-
ples successfully led to the isolation of electroactive microbes, including Ente-
rococcus faecalis, Escherichia coli and Klebsiella pneumoniae, that can possibly
serve as model representatives of the human gut. Isolation of new microbes from
fecal samples motivates further electrochemical characterization of their electron
transfer pathways. This will be important to understand unfamiliar anaerobic
metabolisms in the human gut environment and their implications towards gut
health. Outward electron transfer in E. faecalis depends upon a
avin based EET
mechanism, however the strain in incapable of synthesizing the
avins [186]. This
implies that in the natural gut environment, EET in Enterococcus may rely upon
syntrophic interactions with other
avin producing microbes like E. coli [117] or
Faecalibacterium prausnitzii [91]. Such interactions can also be studied electro-
chemically in bioreactors with co-cultures of the microbes under investigation.
88
In chapter 5, an ecient bacterial immobilization technique was developed that
consistently worked across phylogenetically diverse microbes studied in this thesis,
including S. oneidensis MR-1, P. aeruginosa PA14 and T. electrotropha ElOx9, to
enable AFM studies under liquid droplet. This study encourages AFM character-
ization of cells under physiological conditions. With the immobilization protocol
nalized, specialized conductive probe with insulated cantilever demonstrating low
leakage currents from Scuba Probe Technologies (Alameda, CA), can be used to
perform AFM-SECM. This is a two-pass technique, where the rst pass will deter-
mine the cellular topography and second pass will determine their electrochemical
activity using feedback mode with
avins acting as shuttles for electron transfer
between cells and the probe. Development of this technique will have important
implications in identication and quantication of electroactivity of microbes from
a co-culture of microbes.
89
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Abstract (if available)

Abstract Extracellular electron transfer (EET) is a metabolic process that allows microorganisms to oxidize or reduce solid electron donors or acceptors including insoluble minerals. This phenomenon governs many biogeochemical cycles of major elements including iron, sulfur, manganese, carbon and nitrogen. This process can be mimicked on favorably poised electrode surfaces and has important implications in microbial electrochemical technologies including microbial fuel cells, wastewater treatment, bioremediation and microbial electrosynthesis. The efficiency of these operations relies largely on our understanding of the pathways that dictate EET. ❧ With biophysical, biochemical and genetic toolkit at our disposal, our understanding of EET mainly arises from studies performed on model metal reducing microbes belonging to the genera, Shewanella and Geobacter. Nonetheless, the EET phenomenon is diverse and widespread and many of its aspects are still unclear. (1) Outward EET mechanisms (cell-to-electrode) have been exhaustively characterized, however, the reverse process of inward EET (electrode-to-cell) remains poorly understood, predominantly due to the lack of model organisms and difficulty in laboratory handling of current model microbes. (2) The studies used to explore the scope of EET have been restricted mostly to natural environments, however, anoxic niches of the human body that harbor a vast diversity of microbes may also house novel electroactive microbes that are still uncharacterized. (3) Techniques used for characterizing EET, especially bulk electrochemistry, reports the behavior of a large microbial population but fails to reveal single-cell electron transfer properties or even differentiate between electroactive and non-electroactive cells. ❧ This thesis addresses these issues in three parts. In the first part, we investigate the possible electron uptake mechanisms of a recently isolated marine microbe, Thioclava electrotropha ElOx9. This sulfur oxidizer has been previously shown to respire oxygen using reduced electrodes as a stable carbon source. However, a detailed electrochemical characterization of inward EET pathways is lacking and only limited to aerobic conditions. Here, we use amperometric and voltammetric tools to demonstrate that ElOx9 can couple oxidation of cathode to reduction of nitrate when it is used the sole electron acceptor. Our data reveals that the electron uptake by ElOx9 is facilitated by direct contact, using a redox center with a formal potential of -94 mV vs SHE, instead of soluble redox mediators. ❧ In the second part, we examine EET in host associated bacteria residing in anaerobic regions of human lungs and gut. Pseudomonas aeruginosa PA14, a lung microbe, has been previously shown to respire anodes using phenazines as a soluble redox mediator. However, bioelectrochemical data endorsing this mechanism have been limited. Here, we undertake a thorough electrochemical characterization of PA14 phenazine mutant (Δphz) cells with exogenously added phenazines, thereby confirming that phenazines enable electron shuttling between the cells and an external electrode. We also demonstrate that the efficiency of phenazine redox cycling directly depends upon the extent of mixing in the bioelectrochemical system. Additionally, we attempt to qualitatively determine whether PA14 (Δphz) cells respiring anodes experience any bioenergetic gain, by conjugating amperometric analysis with fluorescence microscopy and using a fluorescent dye Thioavin T, capable of reporting changes in cellular membrane potential. Our results indicate that the cells likely experience a low energetic gain due to low electron transfer rates. ❧ Pertaining to the anaerobic gut microbiome, most resident microbes are known to undergo low energy yielding fermentation. However, other forms of metabolism, including EET, may also be prevalent and remain unexplored. Here, we perform anodic enrichment of facultative microbes from a community outgrown from human fecal samples, representative of the large intestine. Seven morphologically distinct colonies were isolated from electrode attached biofilms, three of which were identifed by 16S rRNA sequencing as Enterococcus faecalis, Klebsiella pneumoniae and Escherichia coli. An increase in anodic current upon inoculation of these isolates in bioreactors indicate possible EET capability of these microbes and are currently under investigation. ❧ In the third part, we focus on determining an efficient and consistent bacterial immobilization technique applicable over diverse range of microbes for atomic force microcopy (AFM) imaging under liquid. Here, we successfully immobilized three phylogenetically diverse electroactive bacteria focused in this dissertation - Shewanella oneidensis MR-1, Thioclava electrotropha ElOx9 and Pseudomonas aeruginosa PA14, on poly-L-lysine functionalized silicon wafers using 5% glutaraldehyde as a crosslinker. Topographical images of the immobilized cells obtained from tapping mode AFM revealed smooth surface structure in liquid as compared to drying related artifacts observed when imaged in air. With the help of fluorescence microscopy using LIVE/DEAD stain, it was observed that most of the immobilized microbes were viable and not affected by the chemical modification. Phase contrast between the substrate and cells, obtained from phase images was taken as a measure of cell adhesion and its quantification revealed that ElOx9 cells were strongly adhesive, followed by MR-1, then PA14 cells. ❧ The work conducted in this thesis has important implications in understanding the functional diversity of EET and the extent of environments where EET-capable microbes may reside. T. electrotropha ElOx9, in addition to serving as a model organism for understanding inward EET mechanism, may also help in understanding biogeochemical sulfur oxidation in marine environments. A thorough mechanistic characterization of gut microbes isolated as a result of electrode enrichment, may further our understanding of unique anaerobic metabolisms in the gut environment that had not been explored yet. Manipulation of EET pathways through biosynthetic techniques can help externally control bacterial population associated with human gut. Additionally, developing sophisticated tools like AFM-SECM will enable detection of electroactive bacteria and simultaneous quantification of their activity from a mixed culture of microbes.

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University of Southern California Dissertations and Theses

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Asset Metadata

Creator Karbelkar, Amruta Anand (author)

Core Title Electrochemical investigations and imaging tools for understanding extracellular electron transfer in phylogenetically diverse bacteria

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Chemistry

Publication Date 12/17/2019

Defense Date 08/30/2019

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag bacterial respiration,extracellular electron transfer,OAI-PMH Harvest

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor El-Naggar, Mohamed (committee chair), Finkel, Steven (committee member), Narayan, Sri (committee member)

Creator Email amrutakarbelkar@gmail.com,karbelka@usc.edu

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c89-257717

Unique identifier UC11673470

Identifier etd-KarbelkarA-8084.pdf (filename),usctheses-c89-257717 (legacy record id)

Legacy Identifier etd-KarbelkarA-8084.pdf

Dmrecord 257717

Document Type Dissertation

Rights Karbelkar, Amruta Anand

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

Access Conditions The author retains rights to his/her dissertation, thesis or other graduate work according to U.S. copyright law. Electronic access is being provided by the USC Libraries in agreement with the a...

Repository Name University of Southern California Digital Library

Repository Location USC Digital Library, University of Southern California, University Park Campus MC 2810, 3434 South Grand Avenue, 2nd Floor, Los Angeles, California 90089-2810, USA