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Extracellular electron transport: Investigating the diversity and mechanisms behind an understudied microbial process with global implications

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Extracellular electron transport: Investigating the diversity and mechanisms behind an understudied microbial process with global implications

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Content EXTRACELLULAR ELECTRON TRANSPORT: INVESTIGATING THE
DIVERSITY AND MECHANISMS BEHIND AN UNDERSTUDIED MICROBIAL
PROCESS WITH GLOBAL IMPLICATIONS

by

Bonita Rasmey Lam

A Dissertation Presented to the
FACULTY OF THE GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(BIOLOGY)

May 2018

Copyright 2018 Bonita Rasmey Lam
ii
DEDICATION

This work is dedicated to my parents who continually inspire me with all that they have done and
all that they do. Their unwavering love and support is instrumental to me being who I am today. I
also dedicate this to my sister, who never fails to make me laugh and has been with me for as
long as I’ve known. I would like to also dedicate this to all my dear family and friends – I hope
you all know how important you are to me.

iii
ACKNOWLEDGEMENTS

I would like to thank the University of Southern California graduate school for supporting me
with a Provost Fellowship to allow me to pursue this research. Multiple Wrigley Institute
Graduate Summer Fellowships were integral in supporting this work and providing the tools and
facilities necessary for sample collection and microcosm construction and incubation. A JPL
Strategic University Partnership Program award supported the work presented in Chapter 4. I
would like to acknowledge everyone I have had the privilege to work with during my time as a
graduate student at USC: my advisor who possesses an enthusiasm for science that is
unparalleled, Dr. Kenneth Nealson, my qualifying and dissertation committee members (Dr. Jan
Amend, Dr. Mohamed El-Naggar, Dr. Steven Finkel, Dr. Karla Heidelberg, Dr. Wiebke Ziebis,
and Dr. Will Berelson), the postdocs who have helped and guided me through this whole process
(Dr. Annette Rowe, Dr. Lina Bird, Dr. Reed Li, Dr. Akihiro Okamoto, Dr. Roman Barco, and
Dr. Nancy Merino), my collaborators at JPL (Dr. Aaron Noell and Dr. Laura Barge), and my
fellow labmates for all their support and assistance: Pratixa Savalia, Casey Barr, Brittnay Hong,
Katie Lee, Stephen Chen, Pournami Rajeev, Rachel Chang, Diana Hang, William Tran, and Dr.
Prithivi Chellamuthu. I would like to thank members of the WMSC staff who made my summer
research possible including Tyler Hild, Lauren Czarnecki Oudin, and Kellie Spafford.

iv
TABLE OF CONTENTS

DEDICATION ............................................................................................................................... ii
ACKNOWLEDGEMENTS ......................................................................................................... iii
LIST OF TABLES ....................................................................................................................... vii
LIST OF FIGURES .................................................................................................................... viii
ABSTRACT ................................................................................................................................... x

CHAPTER 1. Introduction ............................................................................................................. 1
Background .......................................................................................................................... 1
Summarized Questions and Hypotheses ............................................................................. 3
References ........................................................................................................................... 7

CHAPTER 2. Community Structure of Cathode-Oxidizing Microorganisms from Marine
Sediment Capable of Extracellular Electron Transport
Introduction ....................................................................................................................... 10
Materials and Methods
Sediment collection and microcosm construction ................................................. 12
Three-electrode system set-up for electrode enrichment and linear sweep
voltammetry of sediments ..................................................................................... 13
Sediment microprofiles of pH, oxygen, and redox ................................................ 14
Geochemical analyses ........................................................................................... 15
DNA extraction of electrode biofilms ................................................................... 15
16S rRNA sequencing and analysis ...................................................................... 16
Genome predictions using 16S rRNA sequences .................................................. 17
Results
Linear sweep voltammetry of sediment microcosms ............................................ 18
Sediment microcosm depth profiles of pH, oxygen, and redox ............................ 19
Iron species, nitrite, and nitrate measurements ................................................ 20-21
Average current generation with varied electrode potential .................................. 22
Electrodes post-incubation .................................................................................... 23
Species richness indicators and diversity estimates of samples ............................ 24
Microbial community richness and evenness of microcosm communities ........... 25
Relative abundance of top OTUs at phylum, class, and order levels .................... 27
Non-metric multidimensional scaling (NMDS) ordination plots .......................... 29
Ordination based heatmaps of microcosm communities ....................................... 30
Relative abundance of predicted KEGG pathways in microcosm communities ... 33
Abundance of KEGG orthologs related to motility ............................................... 35
Abundance of KEGG orthlogs related to metabolic reductases ............................ 36
Discussion .......................................................................................................................... 38
Conclusion ......................................................................................................................... 43
References ......................................................................................................................... 44

CHAPTER 3. Electrochemical Enrichment and Isolation of Novel Microorganisms Capable of
Cathode Oxidation from Marine Sediment
Introduction ....................................................................................................................... 50
v
Materials and Methods
Enrichment media composition ............................................................................. 52
Secondary enrichment in sediment free bioreactors .............................................. 53
Media exchange and analysis of flavin, proteins, and anion in bioreactors .......... 54
DNA extraction and 16S rRNA community analysis of planktonic and biofilm
communities .......................................................................................................... 54
Genomic predictions based on 16S rRNA sequences ........................................... 56
Tertiary enrichment using insoluble electron donor media and isolation ............. 56
Chronoamperometry and cyclic voltammetry of isolates ...................................... 57
Scanning electron microscopy sample preparation and imaging .......................... 58
Phylogenetic analysis of isolates ........................................................................... 58
Results
Secondary bioreactor general summary run information ...................................... 59
Chronoamperometry of secondary bioreactors ..................................................... 60
Measured flavins, proteins, and anions in secondary bioreactors ......................... 61
SEM images of planktonic cells from secondary bioreactors ............................... 62
Species richness and diversity estimates of secondary bioreactor communities ... 64
Relative abundance of the top OTUs at the class and order levels of the secondary
bioreactor planktonic and biofilm communities .................................................... 65
Non-metric multidimensional scaling (NMDS) ordination plots of secondary
bioreactor communities ......................................................................................... 68
Relative abundance of predicted KEGG pathways in bioreactor communities .... 69
Abundance of KEGG orthologs related to metabolic reductases in bioreactor
communities .......................................................................................................... 71
Relative abundance of the top OTUs at the class and order levels of Fe
0
/SO
4
tertiary enrichment cultures ................................................................................... 73
Maximum likelihood phylogenetic tree of Catalina Isolates ................................. 74
Chronoamperometry tests of electrochemical activity of isolates ......................... 75
Cyclic voltammetry tests of isolates ...................................................................... 77
Isolate electrochemical activity summary and midpoint potentials ....................... 78
Flavin production by Gammaproteobacteria isolates ........................................... 79
SEM images of isolates on carbon felt electrodes ................................................. 80
Discussion .......................................................................................................................... 80
Conclusion ......................................................................................................................... 85
Appendices ........................................................................................................................ 86
References ......................................................................................................................... 87

CHAPTER 4. Detecting Microbial Metabolism and Identifying Biosignatures in Soils with
BioElectroChemical Devices
Introduction ....................................................................................................................... 94
Materials and Methods
Overview of BECD experiments ........................................................................... 98
Soil, anolyte, and catholyte preparation for MFC experiments ............................. 99
Minimal media, cell culturing, and inoculum preparation for three-electrode
chemical cell experiments ................................................................................... 100
Bioelectrochemical devices experimental set-up ................................................ 102
vi
Cyclic voltammetry on test substrates ................................................................. 103
Analysis of organic acids and proteins ................................................................ 103
Scanning electron microscopy (SEM) sample preparation of electrodes ............ 104
Results
Cyclic voltammetry of different test substrates and solutions ............................. 104
Long-term microbial fuel cell experiments ......................................................... 105
Microbial fuel cell experiments with organic acid stimulation ........................... 106
Organic acid concentrations measured during the time course of fuel cell
experiments .......................................................................................................... 108
Total proteins concentrations from the soil anolyte (planktonic) communities .. 110
Total proteins concentrations from the electrode (biofilm) communities ........... 111
Three-electrode electrochemical cell experiments with different initial cell
inoculums ............................................................................................................ 112
Discussion ........................................................................................................................ 113
Conclusion ....................................................................................................................... 117
Appendices ...................................................................................................................... 118
References ....................................................................................................................... 120

CHAPTER 5. Conclusion and Future Perspectives ................................................................... 124
References ....................................................................................................................... 126

BIBLIOGRAPHY ...................................................................................................................... 127

vii

LIST OF TABLES

Table 2.1 Nitrite, ferric iron, and ferrous iron concentrations in sediment microcosms during the
first incubation (March 2014) ........................................................................................................ 20

Table 2.2 Average final nitrate, nitrite, ferric iron and ferrous concentrations in sediment
microcosms during the second incubation run (June 2014) .......................................................... 21

Table 2.3 Species richness and diversity indices of microcosm community samples .................. 24

Table 2.4 Relative abundances of predicted KEGG orthologs assigned to KEGG pathways in
microcosm community samples .................................................................................................... 34

Table 3.1 Summary of secondary bioreactor general run information ......................................... 59

Table 3.2 Measured pH, proteins, flavins and anions during secondary bioreactor media
exchanges ...................................................................................................................................... 61

Table 3.3 Relative abundances of predicted KEGG orthologs assigned to KEGG pathways in
bioreactor community samples found to have significant differences in abundance .................... 69

Table 3.4 Summary of isolate electrochemical activity and midpoint potentials ......................... 78

Table 4.1 Summary of substrates used in BECD experiments ..................................................... 99

Table 4.2 Concentrations of organic acids measured during the course of fuel cell experiments
with lactate additions ................................................................................................................... 109

viii

LIST OF FIGURES

Figure 1.1 Schematic of known mechanisms of EET .................................................................... 1

Figure 2.1 Sediment microcosm incubation set-up ...................................................................... 13

Figure 2.2 Linear sweep voltammetry of sediment microcosms .................................................. 18

Figure 2.3 Geochemical depth profiles of pH, oxygen, and redox ............................................... 19

Figure 2.4 Average current generated at each applied redox potential ........................................ 22

Figure 2.5 Photograph of representative electrodes after incubation ........................................... 23

Figure 2.6 Richness and evenness of microcosms communities .................................................. 25

Figure 2.7 Relative abundance of the top OTUs at the phylum, class, and order levels .............. 27

Figure 2.8 Non-metric multidimensional ordination plots of microcosm communities .............. 29

Figure 2.9 Ordination based heatmaps at the order level of the Proteobacteria phylum ............. 30

Figure 2.10 Ordinated based heatmaps at the genus level for the Deltaproteobacteria and
Alphaproteobacteria classes .......................................................................................................... 32

Figure 2.11 Relative abundance of KEGG orthologs assigned to chemotaxis (K03407) and
flagella (K02418) in each microcosm community ........................................................................ 35

Figure 2.12 Relative abundance of KEGG orthlogs assigned to nitrate reductase (K02570) and
thiosulfate reductase (K08352) in each microcosm community ................................................... 36

Figure 3.1 Electrochemical bioreactor cell design and set-up used for secondary bioreactor
enrichments .................................................................................................................................... 52

Figure 3.2 Chronoamperometry profiles of secondary bioreactors .............................................. 60

Figure 3.3 SEM imagines of planktonic cells from the -400 and -500 mV bioreactors ............... 62

Figure 3.4 Species richness and diversity indices of secondary bioreactor communities ............ 64

Figure 3.5 Relative abundance of the top OTUs at the class and order levels for the secondary
bioreactor communities ................................................................................................................. 65

Figure 3.6 Non-metric multidimensional scaling of a Bray-Curtis dissimilarity matrix of the
secondary bioreactor communities ................................................................................................ 68
ix

Figure 3.7 Relative abundance of KEGG orthologs assigned to adenylylsulfate reductase
(K00394) and nitrate reductase (K00370) in secondary bioreactor communities ......................... 71

Figure 3.8 Relative abundance of the top OTUs at the class and order level for Fe
0
/SO
4
tertiary
enrichment cultures ....................................................................................................................... 73

Figure 3.9 Maximum likelihood phylogenetic tree of Catalina isolates ....................................... 74

Figure 3.10 Representative chronoamperometry of isolates ........................................................ 75

Figure 3.11 Cyclic voltammetry of isolates in planktonic and biofilm phases ............................ 77

Figure 3.12 Flavin production by Gammaproteobacteria isolates ............................................... 79

Figure 3.13 SEM images of isolates on carbon felt electrodes .................................................... 80

Figure 4.1 General schematics of the bioelectrochemical devices used ..................................... 101

Figure 4.2 Cyclic voltammograms of different test substrates ................................................... 105

Figure 4.3 Long-term (2 week) baseline fuel cell tests conducted on both soil mixtures .......... 106

Figure 4.4 Fuel cell experiments with organic acid additions conducted on HF Soil ................ 107

Figure 4.5 Fuel cell experiments with organic acid additions conducted on MS Soil ............... 108

Figure 4.6 Average protein concentrations from soil anolyte samples taken during the time
course of replicate fuel cell with organic acid addition experiments .......................................... 110

Figure 4.7 Average protein concentrations from electrodes harvested after replicate fuel cell
with organic acid addition experiments ....................................................................................... 111

Figure 4.8 Current density generated in three-electrode electrochemical cells inoculated with
differing cell inoculums of S. oneidensis MR-1 .......................................................................... 112

x
ABSTRACT
The influence of microorganisms on our world is tremendous as they are primary drivers of
biogeochemical cycles and regulators of ecosystem processes. Their influence comes from the
diverse and often complex metabolic capabilities that they possess, allowing the utilization of a
wide range of substrates for the derivation of energy. The substrates they can use as either
electron donors or acceptors was once thought to be limited to those that were soluble and could
readily enter the cell membrane. The discovery of extracellular electron transport (EET) has
broadened the diversity of substrates to include insoluble substrates such as minerals and metals.
EET is the ability of microbes to transfer electrons to and from insoluble substrates outside of the
cell. Much of the knowledge about EET has been gained through investigations of model
organisms from two genera, Geobacter and Shewanella, and these studies have mainly focused
on how electrons are transferred from cells to insoluble substrates. The process of receiving
electrons from insoluble substrates has largely remained unexplored, and the mechanisms of
solid substrate oxidation are poorly characterized and understood.

The goal of this dissertation work was to further understanding of EET with a focus on cathode
(insoluble substrate) oxidation. Microbes capable of insoluble substrate oxidation have been
elusive to culturing, in part due to the difficulty in maintaining the conditions they require in the
laboratory. Electrochemical techniques have allowed us to mimic the redox conditions provided
by solid substrates and study the process of insoluble substrate oxidation in a quantifiable and
controllable manner. Combining electrochemical techniques with microbiological and molecular
methods, I was able to investigate the influence of redox potential on cathode-oxidizing
community structure. Overall community diversity and richness increased with more negative
applied redox potentials. In addition, abundances of important known EET groups, including the
xi
Altermonadales, Clostridiales, and Desulfuromonadales, varied with redox potential. Motility
and chemotaxis genes were found in greater abundance in electrode communities, suggesting the
importance of these pathways for colonization and utilization of the electrode as an electron
donor. The cathode-oxidizing community enrichments demonstrated the validity of this approach
in capturing groups that are known to participate in EET and also highlighting potentially
important novel groups (e.g. Campylobacterales) that perform EET as well. The initial
enrichments and the insights gleaned from molecular analyses laid the foundation for the
successful isolation of six new strains of bacteria from five different classes capable of cathode
oxidation. These isolates are not only phylogenetically diverse, but they also display varying
electrochemical properties suggesting different mechanisms of EET. The microorganisms
highlighted in this work will help inform potential genetic markers for future studies as well as
aid in developing a framework for detecting EET capabilities in environmentally relevant
microbes. The potential application of bioelectrochemical devices (BECDs) to probe for
microbial metabolic activity in environmental samples was also explored. The proof of concept
investigation demonstrated the utility of BECDs as a life detection strategy. This work
contributes to the modernization and improvement of life detection techniques focused on
metabolism. The data herein presented reveals the importance of EET as a widespread metabolic
ability and environmental process, which is only starting to be understood with the advances
made in electromicrobiology.
1
CHAPTER 1

INTRODUCTION
Microorganisms are the dominant life forms on Earth, comprising 60% of all biomass (Whitman
et al. 1998). In fact, they make up more than 90% of the biomass of the ocean alone. Their ability
to thrive and survive in a wide range of environments from oligotrophic waters to the deep
subsurface stems from the great metabolic versatility that exists among them. This metabolic
versatility has further expanded with the discovery of extracellular electron transport (EET)
(Gralnick & Newman 2007; Rabaey et al. 2007). EET is the capability of microorganisms to
transfer electrons to and from insoluble substrates outside of the cell. Much of what is known
about EET comes from studies of model metal reducing organisms in the Shewanellaceae and
Geobacteraceae, involving the movement of electrons out of the cell to insoluble metal oxides
(Lovley et al. 2004; Logan 2009).

Figure 1.1. Schematic of the known mechanisms of EET. A) Direct electron transfer through
outermembrane proteins (multiheme cytochromes), B) mediated electron transfer through
shuttles, and C) direct electron transfer through cellular extensions.

The mechanisms of EET that have been characterized (Figure 1.1) include interaction of the
cellular electron transport chain with an insoluble substrate through surface proteins (multiheme
cytochromes) (Myers & Myers 1992; Kim et al. 2006; Clarke et al. 2011), small molecules (e.g.,
2
flavins, humic acids, etc.) that are proposed to shuttle electrons between the cell surface and
substrate (Newman & Kolter 2000; Marsili et al. 2008; von Canstein et al. 2008; Okamoto et al.
2013), and processes that extend into extracellular space (nanowires) (Reguera et al. 2005;
Gorby et al. 2006; Vargas et al. 2013; Pirbadian et al. 2014). The study of EET is a fairly new
and emerging field, and which much that is unknown about both the metabolic versatility (the
diversity of respiratory systems) and the taxonomic extent (number of different microbial taxa)
of organisms that are EET-capable.

Lithotrophs are known to oxidize inorganic and often insoluble substrates. EET may play an
important role for lithotrophs as they often reside in environments where conductive minerals are
abundant and organic carbon inputs are low (i.e., the deep subsurface). Nevertheless, the
mechanisms of solid substrate oxidation are poorly characterized and understood (Emerson et al.
2010). Since the proteins involved in insoluble substrate oxidation remain unknown, they can
potentially be overlooked and undetected in “-omic” centered studies that dominate a lot of
environmental microbiology today. Much of the data generated by “-omic” studies remains
unassigned to any known function, with many genes being grouped together into conserved
hypothetical and/or conserved unknown fractions, and EET genes might comprise some of that
fraction. Our knowledge of the role(s) these microorganisms play in many environments is
extremely limited, in part because these physiologies are difficult to detect in nature solely
through chemistry. This can be attributed to low substrate concentrations, the transient nature of
reactants and/or products, or competition with abiotic or geochemical reactions (Weber et al.
2006). The fact that the conditions lithotrophs require (i.e., maintaining a specific redox
potential) are difficult to replicate in the laboratory, might explain why it has been such a
3
challenge to culture the ‘uncultured majority’ (Jørgensen & Boetius 2007). Microorganism-
electrode interactions are comparable to reactions that occur between microorganisms and
minerals and provide a suitable way to mimic the environmental conditions required for
culturing EET-dependent metabolisms. Electrodes provide a constant (i.e., will not become
soluble once reduced or oxidized) source of substrate that can be poised at a specific redox
potential.

Much of the literature has focused on the anodic side of EET. It has been previously shown that
microorganisms in aquatic sediments are capable of utilizing anodes as electron acceptors, in
doing so generating a current when connected to a cathode in overlying oxygenated waters
(Reimers et al. 2001; Ryckelynck et al. 2005). Enrichment and community characterization of
anode respiring microorganisms has been previously performed in environments such as marine
sediments (Bond et al. 2002; Rabaey et al. 2004; Reimers et al. 2006). Although we know that
microorganisms use electrons from a cathode for reduction of oxygen and different metals
(Rosenbaum et al. 2011), there have been no reports of environmental enrichments of
microorganisms using cathodes until the initial work our laboratory has done (Rowe et al. 2015).

The goals of this dissertation were to build upon this initial work and further explore the use of
electrochemical techniques in combination with microbiological and molecular methods to
investigate the diversity and mechanisms of EET with regards to insoluble substrate oxidation.
The investigations conducted took a multi-faceted approach and were designed to answer the
following questions:

4
Q1: What is the taxonomic diversity of EET-capable microbes (the number of microbial taxa
that can conduct EET)?
Q2: What is the metabolic versatility (the range of respiratory systems) of organisms that utilize
EET?
Q3: Will the application of different cathodic redox potentials in an environmental system enrich
for different microbial groups?
Q4: Will the mechanisms of cathode oxidation be the same as those utilized for anode reduction?
Q5: How prevalent is EET in various environments?

For each of these questions, the following hypotheses were tested in the following chapters.
H1: EET will be found in groups that are not presently known to be electrochemically
active (Chapters 2 & 3).
EET might be common amongst microorganisms present in ecological niches that contain low
organic carbon inputs and an abundance of minerals. These microorganisms have been difficult
to study and culture, but our electrochemical culturing techniques are designed to overcome
these difficulties.
H2: Different redox potentials will enrich for different microbial groups based on
energetics and the optimal terminal electron acceptor used by each group (Chapters 2 & 3).
Electrode-reducing (anodic) communities have been studied in detail in the literature, and it has
been shown that the composition of biofilms change depending on the applied potential and/or
the available carbon source (Nielsen et al. 2009; Sun et al. 2010; Ishii et al. 2012; Ishii et al.
2014). The changes in composition are likely to be a result of the redox potential of the electron
acceptor each group preferentially uses. The changes in community structure in our cathode-
5
oxidizing communities will likely be reflective of the redox potential of the electron donors
available in the environment coupled to the electron acceptors present.
H3: We hypothesize that while some cathode oxidizers will use similar or identical
multiheme cytochromes for EET similar to those in mineral reducers, others will use
different proteins and different pathways and mechanisms (Chapter 3).
Recently the observed homology and partial operon conservation of the MtrAB and MtoAB
genes in the metal reducer, Shewanella oneidensis MR-1 and the iron oxidizer, Sideroxydans
lithotrophicus ES-1 was shown (Liu et al. 2012). This demonstrates a potential conservation in
EET mechanisms between metabolically distinct microorganisms to accomplish different tasks.
Bacterial biochemical pathways mediating sulfur metabolism are well characterized in
facultative sulfur oxidizing Alphaproteobacteria (the Sox system), while very little is known
about the mechanisms of sulfur oxidation in the Beta- and Gammaproteobacteria which
comprise many obligate lithotrophs (Ghosh & Dam 2009). Furthermore, with sulfur oxidation
distinct methods of energy conservation occur for the same inorganic sulfur compound (Finster
2008). The combined approaches of this dissertation work are designed to detect and discover
novel pathways for EET involved with electron uptake.
H4: EET is a relevant process in environments where specific redox niches are conducive to
the energetic yield provided by EET, and electrochemical techniques are a viable way to
identify these processes in the environment (Chapter 4).
Electrochemical techniques can measure electrical potential generated by microbial metabolism
and offer a way to probe for habitable niches in environments traditionally thought to be energy
limited. Environmental samples from habitable niches should possess all the essential
components needed to produce current flow, i.e., catalysts (microorganisms) with fuel and/or an
6
oxidant. In addition, electrochemical techniques can probe for dormant and/or starving
microorganisms by artificially adding a fuel or oxidant to drive metabolism.

7
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9
Rowe, A. R., Chellamuthu, P., Lam, B., Okamoto, A. & Nealson, K. H. Marine sediments
microbes capable of electrode oxidation as a surrogate for lithotrophic insoluble substrate
metabolism. Front. Microbiol. 5, 784 (2015).
Ryckelynck, N., Stecher, H. A. & Reimers, C. E. Understanding the Anodic Mechanism of a
Seafloor Fuel Cell: Interactions Between Geochemistry and Microbial Activity. Biogeochemistry
76, 113–139 (2005).
Sun, Y., Zuo, J., Cui, L., Deng, Q. & Dang, Y. Diversity of microbes and potential
exoelectrogenic bacteria on anode surface in microbial fuel cells. J. Gen. Appl. Micrbiol. 56, 19-
29 (2010).
Vargas, M., Malvankar, N. S., Tremblay, P., Leang, C., Smith, J. A., Patel, P. et al. Aromatic
Amino Acids Required for Pili Conductivity and Long- Range Extracellular Electron Transport
in Geobacter sulfurreducens. mBio 4, 1–6 (2013).
von Canstein, H., Ogawa, J., Shimizu, S. & Lloyd, J. R. Secretion of flavins by Shewanella
species and their role in extracellular electron transfer. Appl. Environ. Microbiol. 74, 615–23
(2008).
Weber, K. a, Achenbach, L. A. & Coates, J. D. Microorganisms pumping iron: anaerobic
microbial iron oxidation and reduction. Nat. Rev. Microbiol. 4, 752–64 (2006).
Whitman, W. B., Coleman, D. C. & Wiebe, W. J. Prokaryotes: The unseen majority. Proc. Natl.
Acad. Sci. U. S. A. 95, 6578–6583 (1998).

10
CHAPTER 2

COMMUNITY STRUCTURE OF CATHODE-OXIDIZING MICROORGANISMS
FROM MARINE SEDIMENT CAPABLE OF EXTRACELLULAR ELECTRON
TRANSPORT

INTRODUCTION
Marine sediments constitute the largest single ecosystem on Earth in spatial coverage (Snelgrove
et al. 2014) and their high conductivity and geochemical complexity provides an excellent
habitat for microorganisms that rely on solid phase minerals for metabolism (Rowe et al. 2015;
Li & Nealson 2015). However, the importance of these metabolisms is not fully elucidated in
these environmental systems as the mechanisms and organisms capable of solid substrate
oxidation remain poorly understood and characterized (Emerson et al. 2010). Various studies
have utilized two-electrode systems to enrich for and characterize anode reducing (metal
reducing) microorganisms in marine sediments (Bond et al. 2002; Tender et al. 2002; Holmes et
al. 2004; Reimers et al. 2006; Mathis et al. 2008). Anodes from marine sediments have been
shown to be enriched in microorganisms in the Geobacteraceae family (Tender et al. 2002; Bond
et al. 2002; Ryckelynck et al. 2005; Reimers et al. 2006) and microbes most closely related to
those in the Desulfobulbaceae family (Tender et al. 2002; Holmes, Bond & Lovley 2004).
Geobacter species have become model organisms for understanding mechanisms of direct
electron transfer to insoluble substrates (Lovley 2012). Within the Desulfobulbaceae family,
Desulfobulbus propionicus has been shown to utilize Fe(III) and graphite electrodes as electron
acceptors (Holmes, Bond & Lovley 2004). In addition, long multicellular cables of
Desulfobulbaceae filamentous cells have been found to transport electrons over centimeter
distances, although the mechanism of electron transport is not yet known (Pfeffer et al. 2012;
11
Risgaard-Petersen et al. 2012; Marzocchi et al. 2014). These studies have expanded our
knowledge about which organisms play a role in anode reduction and are capable of using
insoluble substrates as electron acceptors but the reverse process of cathode oxidation has
remained largely unexplored.

Studies of anodic biofilm communities have demonstrated that different redox potentials and/or
carbon sources will affect the composition of the resultant microbial communities (Finkelstein et
al. 2006; Nielsen et al. 2009; Ishii et al. 2012; Ishii et al. 2014). Different redox potentials were
shown to drive selection and enrichment of distinct microbial communities from sediments that
could optimize the difference in potential energy between microbial terminal electron donors
(e.g. outer-membrane redox proteins) and the anode surface (Finkelstein et al. 2006; Wagner et
al. 2010; Ishii et al. 2014). Selective feeding of specific carbon substrates also impacted the
dominant members of anode microbial communities and consequently the structure of the rest of
the biofilm community (Nielsen et al. 2009; Ishii et al. 2012). Ultimately, understanding the
redox gradients within an environmental system and the oxidative and terminal respiratory
proteins present in endemic microbial communities will suggest the possible metabolisms
present including metabolisms that involve EET processes.

Our laboratory’s pilot study was the first to implement a three-electrode system to study EET
processes in marine sediments, and more specifically it was the first environmental enrichment
of cathode-oxidizing microorganisms (Rowe et al. 2015). Three-electrode systems allow greater
manipulation and control of electrodes to be poised at specific redox potentials versus the
traditional two-electrode systems widely used in the literature. Catalina Harbor (Catalina Island,
CA) was chosen as our source of marine sediment for microcosm construction for electrode
12
incubation due to known high nitrate, ferric, and ferrous iron concentrations (Bertics & Ziebis
2009). These geochemical conditions in addition to low organic carbon inputs compared to other
sites around Catalina Island provide an ideal environment for lithotrophs. The pilot study focused
solely on one redox potential, -400 mV (vs. Ag/AgCl) and no analyses were done on the initial
electrode biofilms enriched from the sediment microcosms.

In this work, a variety of applied redox potentials were chosen between -300 and -750 mV (vs.
Ag/AgCl) to enrich for cathode-oxidizing microbial communities. Using a broad range of redox
potentials allows a greater possibility of capturing different metabolic modes in terms of electron
accepting processes when coupled to cathode oxidation. The applied redox potentials chosen did
not produce any abiotic redox signals, provided adequate energy to predicted terminal electron
accepting processes, and was similar to a range of voltages important for corrosion processes
(Little et al. 1991). In addition, the redox potentials chosen do not cause significant hydrogen
production (Gregory et al. 2004).

We investigated the diversity of microorganisms capable of insoluble substrate oxidation and
shifts in microbial community structure with cathodic potential through combining our newly
developed electrochemical enrichment technique with amplicon based next generation
sequencing. Application of these approaches along with tracking of geochemical gradients and
compounds provides a more complete picture of the potential microbial players and metabolic
processes involved in EET in our cathode-oxidizing communities. These data lead us to suggest
that the two main driving factors for the resulting cathode-oxidizing communities are first, the
redox potential applied to the electrode, and second, the geochemical conditions present around
the electrode.
13
MATERIALS & METHODS
Sediment Collection and Microcosm Construction
Marine sediments were collected from Catalina Harbor (33.4285° N, 118.5090° W) during two
different sampling trips (March, 2014 and July, 2014). Sediments were sieved through 500 µm
copper mesh and transferred to 10-gallon (~40 L) glass aquaria. The sediments were allowed to
settle for 2 weeks before electrochemical enrichment in a constant temperature room maintained
at 15°C. UV-treated, 0.2 µm filtered seawater was pumped over each sediment microcosm at a
rate of ~20 L/day to replenish oxygen and flush out any metabolic waste products.

Figure 2.1. Sediment microcosm incubations. A) Photograph of the sediment microcosms in
the temperature controlled room, and B) schematic of the three-electrode system used with
indium tin oxide working electrodes, Ag/AgCl reference electrode and platinum counter
electrode.

Three-electrode System Set-Up for Electrode Enrichment and Linear Sweep Voltammetry of
Sediments
For electrode enrichment, working electrodes consisted of replicate indium tin oxide (ITO)
plated glass electrodes (SPD Laboratory Inc., Hamamatsu, Japan) buried approximately 3 cm
below the microcosm sediment-water interface. Working electrodes (1.5 cm x 7.5 cm) were
constructed by bonding insulated copper wire to ITO plated glass with Dotite
®

Silver Paint (SPI
Supplies, West Chester, PA). Marine grade epoxy (Loctite, Henkel Co., Rocky Hill, CT) covered
A) B)
14
the connection between the wire and ITO plated glass. Resistance for all electrodes was checked
and only electrodes with a resistance below 30 Ω were used. Coiled platinum wire (Alfa Aesar,
Tewskbury, MA) served as counter electrodes. Reference electrodes (Ag/AgCl, 0.1 M KCl) were
made in our laboratory; all redox potentials reported here are vs. Ag/AgCl. Counter and
reference electrodes were placed in the overlying seawater of each sediment microcosm (see
Figure 2.1B for the three electrode system set-up).

Linear sweep voltammetry (LSV) tests were performed on the sediments using a Pine
potentiostat (Pine Research Instrumentation, Durham, NC) to determine which applied redox
potentials would not result in significant abiotic electron uptake. An ITO electrode was placed at
an approximate depth of 3 cm below the sediment-water interface for LSV tests. LSV tests were
conducted with a sweep rate of 0.3 mV/sec between 0 and -1000 mV (vs. Ag/AgCl).

Working electrode potentials were maintained and controlled using an eDAQ quad channel
potentiostat (eDAQ Inc., Colorado Springs, CO). Applied redox potentials and current
production were monitored and sampled at a rate of 1 sample/min over the incubation period
with the eCorder eCHART software (eDAQ Inc., Colorado Springs, CO).

Sediment Microprofiles of pH, Oxygen, and Redox
Geochemical profiles of pH, oxygen, and redox were examined using Unisense microelectrodes
and micromanipulator (Unisense, Aarhus, Denmark). All microelectrodes were calibrated and
used in accordance to manufacturer’s protocols (Unisense, Aarhus, Denmark). Profiles were
taken from the sediment-water interface to a depth of approximately 5-6 cm with a step size of
100 µm and a sample measurement period of 0.5 seconds. Sediment microprofiles were taken
before electrode incubation and weekly during the course of the enrichment process.
15
Geochemical Analyses
Sediment core samples were taken of the sediment microcosms at three different time points
during the first incubation run and once at the end of the second incubation run for geochemical
analyses. Collected sediment cores were stored at -80°C until porewater extraction. Porewater
was extracted from the sediment core sections through centrifugation at 14,000 rpm for 45
minutes at 4°C. The sum of nitrate and nitrite was determined spectrophotometrically after
reduction of porewater samples with spongy cadmium (Jones 1984). Ferric and ferrous iron
concentrations were determined spectrophotometrically using the Ferrozine assay on wet
sediment and porewater (Stookey 1970). All spectrophotometric measurements were made with
a Shimadzu UV-2600 Spectrophotometer (Shimadzu Corporation, Kyoto, Japan).

DNA Extraction of Electrode Biofilms
Electrodes recovered after sediment microcosm incubation were immediately frozen and kept at
-80°C until further processing. For DNA extraction, biofilm from the electrodes was scraped off
and electrodes were treated with sonication in sterile artificial seawater base to remove as much
biomass as possible. The artificial seawater base medium contained 342 mM NaCl, 14.8 mM
MgCl
2
⋅6H
2
O, 0.1 mM CaCl
2
⋅2H
2
O, and 6.7 mM KCl. The electrodes were also treated with 200
µl of lysozyme and 200 µl proteinase-K. The biomass removed from the electrode and the 400 µl
of enzymatic treatment was directly added to bead beating tubes from a MO BIO PowerSoil
DNA Isolation Kit (Qiagen, Carlsbad, CA). DNA extraction was performed using the MO BIO
Powersoil DNA Isolation Kit according to manufacturer’s protocols. Extracted DNA quality and
quantity was determined using a Nanodrop (Thermo Fisher Scientific, Wilmington, DE).

16
16S ribosomal RNA (rRNA) Sequencing and Analysis
Illumina MiSeq paired-end (2 x 250 bp) sequencing was performed on the V4 hypervariable
region of the 16S rRNA gene of extracted DNA by MR-DNA (Molecular Research LP,
Shallowater, TX). In summary, ribosomal sequences were amplified with a 30-cycle PCR
reaction using the HotStarTaq Plus Master Mix Kit (Qiagen, Carlsbad, CA) with a 53°C
annealing temp using barcoded 515F (5’-GTG CCA GCM GCC GCG GTA A) primer paired
with 806R (5’-GGA CTA CHV GGG TWT CTA AT) primer. Amplicons were pooled and
purified with calibrated Agencourt Ampure XP beads (Agencourt Bioscience Corporation,
Beverly, MA). Purified PCR product was used to prepare DNA libraries by following the
Illumina TruSeq DNA library preparation protocol and sequenced on an Illumina MiSeq
following manufacturer’s guidelines (Illumina Inc., San Diego, CA). Average sequence lengths
of 250 bp and > 35,000 sequences were obtained for all samples.

Quantitative Insights Into Microbial Ecology (QIIME v. 1.9.1) bioinformatics pipeline (Caporaso
et al. 2010) was utilized for quality filtering (Bokulich et al. 2013), chimera checks (Edgar et al.
2011), and barcode removal. De novo operational taxonomic unit (OTU) picking was used for
taxonomy assignment (based on 97% similarity), sequence alignment (Caporaso et al. 2010), and
phylogenetic analysis using the Greengenes database (Werner et al. 2012). The R package,
Phyloseq (McMurdie & Holmes 2013) was implemented for downstream analysis of 16S rRNA
sequencing data including species richness and diversity indices, non-metric multidimensional
scaling (NMDS) and heatmap ordination plots. To avoid artifacts of sequencing depth, sequence
data were normalized to the sample with the lowest sequence yield (n = 38,964 sequences).
Normalized data were then used to calculate diversity indices, NMDS plots, relative abundance
charts, and heatmaps. Permutational multivariate analysis of variance (PERMANOVA) was used
17
to evaluate the contribution of different variables to microbial community variation with the
Adonis function (Anderson 2001). Heatmaps were constructed with different dimension-
reduction algorithms on subsets of community data for ordering over hierarchical analysis to
better preserve the integrity of the topology of the data (Rajaram & Oono 2010). Other statistical
tests including analysis of variance (ANOVA) and Tukey honest significant difference (HSD)
tests were conducted in the R program version 3.4.1 (R Project Team, http://www.r-project.org).
The raw sequences have been uploaded to the NCBI Sequence Read Archive (SRA) database
(accession number: SRP124740).

Genomic Predictions Using 16S rRNA Sequences
Functional composition of the microbial communities from each sample was predicted using the
Phylogenetic Investigation of Communities by Reconstruction of Unobserved States (PICRUSt
v. 1.1.2) bioinformatics software package (Langille et al. 2013). Gene family estimates were
based on an OTU table generated from 16S rRNA sequences processed by closed-reference OTU
picking against the Greengenes database through QIIME (Caporaso et al. 2010). The OTU table
was rarefied to the same number of sequences per sample (n = 26,540). Functional predictions
were based on KEGG (Kyoto Encyclopedia of Genes and Genomes) orthologs (KO) at three
hierarchical levels, clusters of orthologous groups (COG), and RNA families (Rfam).
Statistically significant differences between KO abundances in different sample groups (redox
potentials) were determined using nonparametric Kruskal-Wallis tests.

RESULTS
Linear Sweep Voltammetry: To determine the potential influence of background abiotic
electrochemical reactions, a slow electrode sweep technique, linear sweep voltammetry was
18
employed. Initial linear sweep voltammetry tests on sediment microcosms (Figure 2.2) indicated
that the range of redox potentials (-300 to -750 mV vs. Ag/AgCl) chosen for these cathode-
oxidizing enrichments fell within a range of applied potentials where minimal background
geochemical reactions occur in this sediment. This was indicated by the sub-nA currents
observed in this range and a near 100 mV buffer as no substantial abiotic cathodic current was
generated until about -850 mV (vs. Ag/AgCl). No significant anodic current was observed until
about 0 mV (vs. Ag/AgCl).

Figure 2.2. Linear sweep voltammograms. Results of linear sweep voltammetry tests
conducted with a scan rate of 0.3 mV/sec taken on sediment microcosms before electrode
incubation. Tests indicate no substantial abiotic current is generated in the redox potential range
of interest between -300 and -750 mV vs. Ag/AgCl.

Geochemical Profiles and Analyses: Investigation of the geochemical reactions that could
support the electrode processes of cathode oxidation were evaluated by measuring the
19
concentrations of various geochemical compounds in the sediment and seeing how pH, redox,
and oxygen might change in response to the introduction of an electron donor source (electrode).
During the course of electrode incubation, geochemical depth profiles of pH, oxygen, and redox
potential exhibited consistent patterns. A representative set of geochemical depth profiles is
shown in Figure 2.3. The pH along the depth gradient stayed within a normal range of 7.0-7.5.
Oxygen was rapidly consumed in all sediment microcosms and could not be detected below the
first centimeter. Redox potential depth profiles tracked well with the availability of oxygen,
becoming more reducing with depth. The electrodes were buried at approximately 3 cm where
no oxygen was detected, indicating that coupling of oxygen reduction with electrode oxidation

Figure 2.3. Geochemical depth profiles. Microprofile depth diagrams of A) pH, B) oxygen,
and c) redox potential in each sediment microcosm from the March 2014 incubation run
constructed from 0 cm (sediment-water interface) to 6 cm. Dashed lines indicate where
electrodes were incubated at approximately 3 cm.

20
was not likely and anaerobic processes predominated in the cathode-oxidizing communities.
Other potential electron acceptors in the form of nitrate and Fe(III) were also measured in the
sediment microcosms. Due to limitations in sediment core and porewater volume, nitrate and
nitrite were only measured in samples from one aquarium (Aquarium 3) from the first incubation
run. Table 2.1 summarizes the Fe(II) and Fe(III) concentrations measured in wet sediment and
the NO
2
measured in porewater during the course of the incubation run. Generally, Fe(II)
concentrations were comparable to measurements made in a previous study of Catalina Harbor
sediments (Bertics & Ziebis 2009), falling between 0.028-1.29 mM. Fe(III) was only detected in
small quantities (0.014 and 0.072 mM) at the middle sampling date in two aquaria. NO
2
concentrations declined from 74.22 to 5.60 µM during the course of the experiment for the one
aquarium where measurements were possible. NO
3
could not be measured in the same samples,
suggesting nitrate was rapidly used up and no longer present with time. For the second

Table 2.1. Nitrite, ferric iron, and ferrous iron concentrations in sediment microcosm cores
sectioned for the 3 cm depth during the first incubation (March 2014) at three different
time points

ND indicates not detected
NM indicates not measured
Time points correspond to the following dates: T
1
(4/30/14), T
2
(5/10/14), T
3
(5/27/14)
Incubated electrode redox potentials in each aquarium indicated in parentheses vs. Ag/AgCl

incubation, triplicate core samples were collected at the end of the experiment. Table 2.2
summarizes the concentrations of Fe(II) and Fe(III) found in the separate fractions of sediment
21
and porewater after centrifugation, and the NO
2
and NO
3
measured in porewater. Compared to
the first incubation time point, NO
2
concentrations remained similar while NO
3
was found in
appreciable amounts (145.39 to 250.65 µM). Fe(II) concentrations were generally much higher
in the second incubation compared to those seen in the first incubation, while Fe(III) (0.062 to
0.24 mM) was detected across all porewater samples. The differences in concentrations between
different incubations might be linked to seasonal cycling (spring vs. summer). The presence of
NO
3
and Fe(III) during the second incubation suggests that they could potentially serve as
electron acceptors coupled to cathode oxidation.

Table 2.2. Average final nitrate, nitrite, ferric iron and ferrous iron concentrations in
sediment microcosms during the second incubation (June 2014)

Incubated electrodes in each aquarium indicated in parentheses vs. Ag/AgCl

Current Production in Sediment Microcosm Enrichments: At the end of electrode incubation
and enrichment, average current generated at each redox potential was calculated from the
current values recorded every minute during the course of incubations. Negative current is an
indication of electron uptake from the electrodes. Generally negative current increased with more
negative redox potentials, and this trend is especially strong looking across the first incubation
series (Figure 2.4). The -750 mV (vs. Ag/AgCl) redox potential from the second microcosm run
obtained much more negative current compared to values achieved during the first incubation.
22
The two other redox potentials tested during the second incubation, -550 and -650 mV (vs.
Ag/AgCl), exhibited electron uptake much later in the incubation run, never reaching the level of
negative current observed in prior incubations. Seasonal variations between summer and spring
linked to differences in temperature and nutrient cycling could explain why only the most
negative redox potential from the second run was able to generate appreciable amounts of
negative current. The four redox potentials (-400, -500, -600, and -750 mV vs. Ag/AgCl) that
showed considerable amounts of negative current had comparable or significantly greater values

Figure 2.4. Average current generated at each applied redox potential. Current was averaged
over the course of the incubation runs. The first run consisted of four different redox potentials:
-300, -400, -500, and -600 mV (vs. Ag/AgCl). The second run consisted of three different redox
potentials: -550, -650, and -750 mV (vs. Ag/AgCl).
Run 1: March 2014 Run 2: June 2014
23
compared to our initial study where the highest average current achieved was -1.88 µA (log-
modulus transformed value) (Rowe et al. 2015). The electrodes recovered from the sediment
microcosms showed varying degrees of visible growth and precipitate formation (Figure 2.5).
The -750 mV (vs. Ag/AgCl) redox potential that displayed the highest levels of electron uptake,
yielded electrodes with the thickest visible enrichment (Figure 2.5D). Open circuit controls
showed no visible biofilm formation or staining with crystal violet. DNA extraction performed
on the open circuit electrodes resulted in unquantifiable DNA concentrations by Nanodrop and
were not sent for 16S rRNA sequencing.

Figure 2.5. Photograph of representative electrodes. A) Example of an ITO electrode pre-
incubation, B) -300 mV electrode post-incubation and post-extraction, C) -600 mV electrode
post-incubation and post-extraction, and D) -750 mV electrode post-incubation and before
extraction.

16S rRNA Community Analysis: The effect of applying electrochemical stimuli on sediment
microbial communities was investigated by studying shifts in diversity and composition of 16S
rRNA genes. Amplicon next generation sequencing of the 16S rRNA genes from the different
electrodes and sediment samples resulted in variation in the resulting amount of sequences and
observed OTUs obtained. Sediment samples were taken at the start (AQ3A and AQ4A) and end
of electrode incubation (AQ3B and AQ4B). With the exception of the -650 mV (vs. Ag/AgCl)
A) B) C) D)
24
electrode, all electrode samples had a total number of observed OTUs higher than the control
sediment samples. Table 2.3 summarizes species richness and diversity indices of all microcosm
communities. The species richness estimators, ACE and Chao1, both assess that richness
increases with the more negative set redox potentials. The ACE and Chao1 estimators also both
estimate that the bacterial communities of the electrode samples are statistically different from
those of the surrounding sediment in overall species richness. For both incubation runs,
community samples displayed higher diversity as redox potential becomes more negative
determined by increasing values in the Shannon diversity index. The Simpson diversity index
weighs towards the abundance of the most common species when estimating diversity, taking

Table 2.3. Species richness indices and diversity indices of microcosm samples. Electrode
potentials expressed in values vs. Ag/AgCl.

Replicates from the same redox potential are indicated by A and B; sed refers to sediment sample.
1
Denotes samples from the first incubation run
2
Denotes samples from the second incubation run
*
Denotes sequence data to which other samples were normalized

25
into account evenness more heavily. All samples demonstrated high values in the Simpson
diversity index, suggesting high diversity and evenness across conditions without communities
being dominated by a select few species. To further account for differences in sequencing depth,
normalization was applied by rarefying each sample to 1000 reads 100 times through
bootstrapping. Richness and evenness counts were averaged across all 100 trials and can be seen
in Figure 2.6. Figure 2.6A demonstrates a general trend towards greater richness with lower set
negative redox potentials, corroborating the results of the richness indices used in Table 2.3.
Evenness (Figure 2.6B) also followed the same general trend as richness. The -650 mV (vs.
Ag/AgCl) redox potential sample again stands out amongst the electrode samples, deviating from
the trends observed with relatively low richness and evenness numbers comparable to those

Figure 2.6. Average community richness and evenness. A) Richness and B) evenness of each
microcosm community after rarefying each community sample by bootstrapping to 1000 reads
100 times. Replicates from the same redox potential are indicated by A and B; sed refers to
sediment sample.

26
obtained at the -300 mV (vs. Ag/AgCl) redox set potentials. There was a statistically significant
difference between samples based on grouping (applied redox potential and control sediment) in
both richness (F(7,8) = 18.62, P = 0.00023) and evenness (F(7,8) = 17.21, P = 0.0031) as
determined by one-way ANOVA tests.

Relative abundances at the phylum, class, and order level were examined for each of the
microcosm community samples (Figure 2.7). The top 100 OTUs were evaluated for phylum level
differences, representing 55% of all sequences. All samples were dominated by the
Proteobacteria phyla, ranging from 58.8%-83.9% of the total community. All electrode samples
showed a distinct enrichment of Firmicutes, comprising up to 2.2%-28.7% of the microbial
communities compared to only 0.15%-0.67% in sediment samples. Firmicutes have been shown
to dominate communities of thermophilic anodic communities derived from anaerobic sludge
(Wrighton et al. 2008) and marine sediment (Mathis et al. 2008). Furthermore, a Firmicute strain
exhibited EET ability in reducing various insoluble electron acceptors (Wrighton et al. 2011).
The Verrucomicrobia were also enriched in most of the electrode samples. The Verrucomicrobia
are ubiquitous at low abundances in marine environments (Freitas et al. 2012), and have the
highest overall relative abundance in low productivity environments (Yilmaz et al. 2016). In
addition, the Tenericutes were enriched (1.3%-7.2% of the total community) on electrodes set at
the redox potentials (-500, -600, and -750 mV vs. Ag/AgCl) that demonstrated the highest levels
of negative current. Tenericutes are known to be found in association with eukaryotic hosts, but
recently genomes recovered from marine methane seep sediment suggest they may be non-host-
associated, free-living microorganisms (Skennerton et al. 2016).

27
More distinct differences between the electrode and control sediment samples emerge at the class
level. Across all electrode samples there are enrichments of the Alphaproteobacteria, Clostridia,
and Deltaproteobacteria classes. The -650 mV (vs. Ag/AgCl) electrode community deviates
from all samples in microbial community composition with the only enrichment of the OM190
class. The OM190 is an uncultured class of the Planctomycetes that has been shown to be
members of biofilms found on macroalgae (Bengtsson & Øvreås 2010; Lage & Bondoso 2014).

Figure 2.7. Relative abundances for each community sample at the A) Phylum, B) class,
and C) order levels. The top 100 OTUs were evaluated for relative abundance at the phylum
and class level. The top 50 OTUs were assessed for relative abundance at the order level.
Replicates from the same redox potential are indicated by A and B; sed refers to sediment
sample.

28
In general, the Planctomycetes display a preference for a biofilm lifestyle in marine
environments, attaching to surfaces in marine sediments (Gray & Herwig 1996) and marine snow
(Delong et al. 1993).

Going deeper to the order level, increases in specific orders in the Deltaproteobacteria become
apparent. The Desulfuromonadales, Desulfovibrionales, and Desulfobacterales were all present
in higher abundances in the electrode samples. These orders of Deltaproteobacteria are
distinguished by their ability to use sulfate as an electron acceptor, and the Desulfuromonadales
contains members capable of anaerobic respiration with a broad range of compounds including
sulfur, Fe(III), Mn(IV), and nitrate (Garrity et al. 2005). There was also a notable enrichment of
the Rhodobacterales in the electrode samples from the first incubation run. Species within the
Rhodobacterales are capable of sulfur and iron oxidation (Brinkhoff et al. 2008; Straub et al.
1999) and are dominant members of iron corroding biofilms (Dang et al. 2011). The Clostridales
comprise a significant proportion of the communities for all electrode samples, except for the -
650 mV (vs. Ag/AgCl) electrode community. In our laboratory’s prior studies of Catalina
Harbor, Clostridia comprised a significant amount of the communities in tertiary solid substrate
oxidizing enrichments (Rowe et al. 2015) and in anodic electrochemical enrichments (Li &
Nealson 2015). An enrichment of the orders agg27 and 34P16 distinguishes the -650 mV (vs.
Ag/AgCl) electrode from all other samples. Both orders are uncultivated and poorly described.
The -750 mV (vs. Ag/AgCl) electrodes, which exhibited the highest negative current during
electrode incubation, exhibited the highest enrichment of Alteromonadales. Members of the
Alteromonadales have demonstrated EET ability with the reduction of manganese and iron
oxides in pure culture (Myers & Nealson 1988, Lovley et al. 2004).
29
Community Membership Structure: Bray-Curtis and Unifrac dissimilarity indices were used to
quantify hierarchical clustering of community membership structure similarity (Figure 2.8). Both
dissimilarity measures showed distinct clustering based on either redox potential or incubation
runs. The Bray-Curtis dissimilarity index (Figure 2.8A) demonstrates that the similarity between
communities diverged with each successive change in redox potential. Generally, communities
of the same sample type (applied redox potential or sediment) cluster most tightly with each
other. With both dissimilarity indices, the -650 mV (vs. Ag/AgCl) microcosm community

Figure 2.8. Non-metric multidimensional scaling plots of two dissimilarity measures A)
Bray-Curtis and B) Unifrac applied to the different microcosm communities. Communities
of the same sample type (applied redox potential or sediment) are connected with lines. Arrows
indicate associated trends in the ordering of communities. Replicates from the same redox
potential are indicated by A and B; sed refers to sediment sample.

30
deviated and did not show any great similarity to any of the other samples. Additionally, both
indices show no overlap between control sediment samples and the electrode communities. The
Unifrac dissimilarity index (Figure 2.8B) illustrates clustering based on incubation run.
Communities from the first incubation run (-300, -400, -500, and -600 mV vs. Ag/AgCl) cluster
together while two of the redox potentials from the second incubation run display greater
similarity to one another (-550 and -750 mV vs. Ag/AgCl). Adonis tests were performed on
weighted Unifrac values to determine the significance of variation explained by redox potential,
run (spring vs. summer), and success of incubation as determined by current generation. All of

Figure 2.9. Ordination based heatmaps using two methods A) Detrended Correspondence
Analysis (DCA) and B) Multidimensional Scaling/Principal Coordinates Analysis on the
Unweighted-UniFrac distance. Heatmaps were based on the top 100 orders represented within
the Proteobacteria phylum. Enriched Desulfobacterales (Des) OTUs are indicated in blue,
enriched OTUs (Alteromonadales (Alt), Rhodobacterales (Rho), Rhizobiales (Rhi),
Myxococcales (Myx)) specific to the -650 mV (vs. Ag/AgCl) electrode indicated in yellow.
31
the factors tested significantly impacted microbial community structure (p < 0.01). The largest
source of variation was redox potential (R = 0.91, p = 0.001), followed by current production (R
= 0.72, p = 0.001). Incubation season (spring vs. summer) also influenced variation in microbial
community structure (R = 0.65, p = 0.001).

Heatmap Analyses of Community Structure: Since the Proteobacteria was the most dominant
phylum in all samples, ordination based heatmaps were generated to analyze the finer differences
and clustering between microcosm communities (Figure 2.9). Generally, each sample clustered
within their sample types for both ordination methods, although the ordering of sample types
changed between them. The -650 mV (vs. Ag/AgCl) electrode community had a high abundance
of specific OTUs within the Alteromonadales (Alt), Rhizobiales (Rhi), Rhodobacterales (Rho),
and Myxococcales (Myx) that was not found in the other samples. OTUs within the
Desulfobacterales (Des) were highly enriched in all electrode samples, and were amongst the
few OTUs that were enriched across all redox potentials.

Finer differences were refined at the genus level of the Deltaproteobacteria and
Alphaproteobaceria (Figure 2.10). These two classes showed the greatest enrichment in
electrode communities (Figure 2.7B). Within the Deltaproteobacteria (Figure 2.10A), the
Desulfofaba, Desulfuromonas, and Desulfocapsa were found in abundances up to three orders of
magnitude higher in electrode samples. More specifically, the Desulfofaba and Desulfuromonas
were more highly enriched in the less negative redox potentials (-300 to -600 mV vs. Ag/AgCl),
while the Desulfocapsa were associated with the -650 mV (vs. Ag/AgCl) redox potential. The
Desulfuromonas genus is known to convert elemental sulfur into sulfide and forms a distinct
phylogenetic cluster within the Geobacteraceae (Holmes et al. 2004). Members of the
32
Desulfocapsa are capable of disproportionation of elemental sulfur and thiosulfate (Finster
2008). The -650 mV (vs. Ag/AgCl) electrode community also had the greatest abundance of
Plesiocystis. There were also notable increases in the Pelobacter, Desulfobulbus, and
Desulfobacter genera in the electrode samples. Desulfococcus and Desulfosarcina populations
declined in electrode communities compared to sediment communities.

Looking at the Alphaproteobacteria (Figure 2.10B), some of the highest enrichments occurred
with the Devosia and Octadecabacter found in the -650 mV (vs. Ag/AgCl) electrode community,
while the Marivita were found in the highest abundance (e.g., two orders of magnitude higher) in

Figure 2.10. Ordination based heatmaps looking at the A) Deltaproteobacteria and B)
Alphaproteobacteria classes based on multidimensional scaling/principal coordinates
analysis on the weighted-UniFrac distance. Heatmaps demonstrate changes in abundances at
the genus level within each class.

33
the -300 mV (vs. Ag/AgCl) redox potential communities. There were also increases in the
Phaeobacter for most electrode samples except for the -400 and -500 mV (vs. Ag/AgCl) redox
potentials. Phaeobacter species are known to exhibit a surface-associated lifestyle with
biosynthesis of extracellular polysaccharides, with some strains excreting siderophores under
iron limitation (Thole et al. 2012). Similarities between different samples are also apparent upon
observation of the two heatmaps. When only taking into consideration the Deltaproteobacteria
(Figure 2.10A) there is a separation between set redox potentials. The more negative redox
potentials cluster together (-500 to -750 mV vs. Ag/AgCl), while the other redox potentials form
another cluster (-300 and -400 mV vs. Ag/AgCl). The two more negative redox potentials that do
not group with the rest, -550 and -650 mV (vs. Ag/AgCl), were the ones that did not produce
much negative current during the second incubation run. On the other hand, with the
Alphaproteobacteria (Figure 2.10B) the -300, -600, and -650 mV (vs. Ag/AgCl) redox potentials
show greater similarity. This is substantiated by the fact that those electrode communities had the
highest enrichment in Alphaproteobacteria (Figure 2.7B). The applied redox potential and the
success of the enrichment based on electron uptake rates are determining factors on community
structure.

Predicted genomic content and functionality: PICRUSt calculates the phylogenetic distance
between predicted metagenomes and reference genomes as a way to evaluate the validity of the
metagenome predictions (Langille et al. 2013). Nearest sequenced taxon index (NSTI) values are
calculated to evaluate accuracy, with smaller NSTI values reflecting greater accuracy of
prediction. NSTI values for microcosm community samples ranged from 0.12 to 0.17 for
electrode communities. The NSTI values obtained for electrode community samples fall within
the range reported for accurate metagenome predictions with NSTI values ≤ 0.17 (Langille et al.
34
2013). The sediment microbial communities had slightly higher NSTI values (0.19 to 0.20), and
the predictions for these samples do not capture as much of the potential function present. The
relative abundances of various KEGG pathways related to metabolism, motility and cellular
processes and signaling are reported in Table 2.4. Nitrogen metabolic pathways, carbon fixation
pathways in prokaryotes, and citrate cycle pathways were found in greater abundance in
electrode microbial communities as compared to sediment communities.

Table 2.4. Relative abundances in percentages of predicted KEGG orthologs assigned to
KEGG pathways with relevant functions in the microcosm microbial communities. As a
proxy, values for all KEGG pathways ranged from 0 to 4.58%.

*Indicates a significant difference (p < 0.05) in the abundance of a KEGG pathway grouping with
redox potential by Kruskal-Wallis tests

All motility pathways (bacterial chemotaxis, bacterial motility proteins, and flagellar assembly)
were increased in the electrode communities as well.

The direct contributions to KEGG orthologs (KOs) by specific OTUs were assessed at the order
level with abundance being the total amount of KOs predicted in each sample. Since motility
pathways were found in greater abundance in electrode samples, contributions to chemotaxis and
35

Figure 2.11. Abundance of KEGG Orthologs related to A) Chemotaxis (K03407, two-
component system, sensor kinase CheA) and B) flagella (K02418, flagellar protein
FliO/FliZ) in each microbial community contributed by OTUs at the order level.
36

Figure 2.12. Abundance of KEGG Orthologs related to A) Nitrate reductase (K02570,
periplasmic nitrate reductase NapE) and B) thiosulfate reductase (K08352, thiosulfate
reductase) in each microbial community contributed by OTUs at the order level.
37
flagella were analyzed (Figure 2.11). Both chemotaxis (K03407) and flagella (K02418) KOs
were highest in the electrode communities from the first microcosm run (-300, -400, -500, and
-600 mV vs. Ag/AgCl0). The orders contributing the most to both chemotaxis and flagella KOs
were the Desulfobacterales, Desulfovibrionales, and Desulfuromonadales. The Rhodobacterales
were important contributors to the -650 mV (vs. Ag/AgCl) electrode community with both
motility KOs. The Xanthomonadales factored more into flagella KO (Figure 2.11B) with both
the -300 and -600 mV (vs. Ag/AgCl) electrode communities. Generally, the Thiotrichales had a
bigger impact on flagella KO abundance across all samples. The Altermonadales contribution
was observed in higher abundances in most of the electrode microbial communities for K02418.
K02570, nitrate reductase and K08352, thiosulfate reductase were both found to have significant
differences (P = 0.041 and P = 0.045 respectively) in abundance with redox potential through
Kruskal Wallis tests. Both processes could be important electron accepting reactions when
coupled to electrode oxidation. Figure 2.12 demonstrates that these KOs were found in greater
abundance in electrode microcosm communities and especially with those from the first
microcosm run (-300, -400, -500, and -600 mV vs. Ag/AgCl). The Desulfobacterales group was
the dominant contributor to both K02570 (nitrate reductase) and K08352 (thiosulfate reductase).
The Altermonadales was also another order that was found to contribute to K02570 in all
electrode microbial communities. The Campylobacterales were important for nitrate reductase in
the -500 and -600 mV (vs. Ag/AgCl) microbial communities. The second microcosm run
electrode communities had other orders that were part of the K02570 abundance, including the
Rhodobacterales for the -750 mV (vs. Ag/AgCl) redox potential and the Oceanospirillales and
the Rhodospirillales for the -650 mV (vs. Ag/AgCl) redox potential. There was less variation in
the OTUs associated with K08352. Besides the Desulfobacterales, the Desulfovibrionales and
38
Desulfuromonadales were the main orders influencing the presence of thiosulfate reductase. In
addition, the Syntrophobacterales were a part of the K08352 abundance in the -400, -500, -600,
and -750 mV (vs. Ag/AgCl) electrode communities.

DISCUSSION

Our study utilized a suite of electron donating redox potentials to enrich for microorganisms
capable of insoluble substrate oxidation. The sediment we chose to enrich from has been well
characterized geochemically, demonstrating a wide variety of redox conditions caused by the
disruption of redox gradients by invertebrate burrowing, allowing oxygen to penetrate at deeper
depths (Bertics & Ziebis 2009). To simplify the system for microcosm electrode incubation, the
sediment was sieved to eliminate any burrowing activity by invertebrates. Sieving allowed for
less variation in the redox gradients formed in the microcosm sediment while preserving the
microbial diversity in terms of the possible metabolic potential present in the system. Our initial
study focused on one redox potential, -400 mV (vs. Ag/AgCl) and subsequent enrichment steps
yielded several electrochemically active isolates from the Gammaproteobacteria and
Alphaproteobacteria (Rowe et al. 2015). Many of the strains belong to genera not known to use
insoluble substrates as electron donors, and each strain when characterized electrochemically
demonstrated unique interactions with the electrode indicating variations in the potential EET
mechanisms employed (Rowe et al. 2015). For our current study, a greater range of redox
potentials was chosen to enrich for cathode-oxidizing microorganisms. Applying more reducing
(more negative) redox potentials increases the possibility of enriching for a greater diversity of
microbes. Although our initial study yielded many electrochemical isolates they were limited to
two classes within the Proteobacteria, the Gammaproteobacteria and Alphaproteobacteria.
Various strains of Deltaproteobacteria have demonstrated cathode-oxidizing ability, although
39
they were initially isolated for their mineral reducing capability (Gregory et al. 2004; Dumas &
Bergel 2008; Rosenbaum et al. 2011). Other classes within the Proteobacteria known to have
lithotrophic members are the Zetaproteobacteria and the Epsilonproteobacteria. The
Zetaproteobacteria are a recently described class of iron-oxidizing bacteria (Emerson et al.
2007) that are present in various iron rich environments (Emerson et al. 2010; Hedrich et al.
2011; Chan et al. 2016). The Epsilonproteobacteria are recognized as important members in
hydrothermal vent environments, capable of using reduced sulfur compounds and hydrogen for
energy (Takai et al. 2005; Grote et al. 2008; Waite et al. 2017). These other classes of
Proteobacteria could contribute to the increasing diversity of microorganisms capable of cathode
oxidation and can potentially be enriched by using a broader range of redox potentials.

Many of the redox potentials in our study produced a robust response in terms of electron uptake
activity in the sediment microcosms, ranging from -0.66 to -2.63 µA (log-modulus transformed
values). In our previous work, the -400 mV (vs. Ag/AgCl) redox potential had an average current
of -1.88 µA. Although the -750 mV (vs. Ag/AgCl) redox potential from our second microcosm
run produced the highest observed negative current, the other two redox potentials did not have
as much electron uptake activity. Seasonal differences in the amount of organic carbon loading
into Catalina Harbor might account for less reducing redox potentials not generating much
negative current. During the summer the area around the harbor is impacted by much more
anthropogenic activity. In addition, the concentrations of Fe(II) in the sediments were much
higher in the summer than in the spring. Greater competition from the availability of other
potential electron donors during the summer months might explain why only the most reducing
redox potential was thermodynamically favorable for enrichment.

40
Insights into the geochemistry of the microcosm sediments allowed us to understand the
potential terminal electron accepting processes being used by the cathode-oxidizing
communities. Redox and oxygen geochemical depth profiles demonstrated that oxygen is not a
likely electron acceptor being utilized. Measurements of nitrate and nitrite indicate that nitrate
might be an important electron acceptor for the second microcosm run communities. Declining
concentrations of nitrite in the first microcosm run suggest that nitrate might have become
limited and was the predominant electron acceptor utilized during the course of incubation. No
exogenous sources of nutrients were introduced to the sediment microcosms during the course of
incubation besides flowing filtered seawater. The naturally high levels of sulfate (28 mM)
present in seawater make it a readily available electron acceptor. The higher abundance of
Deltaproteobacteria found in the first microcosm run compared to the second microcosm run
suggest that sulfate was likely an important electron acceptor for the first microcosm’s cathode-
oxidizing communities.

The overall richness of the enriched cathode-oxidizing microbial communities increased with
more negative redox potentials. Dissimilarity indices were used to evaluate the differences in
community structure between the different samples (Figure 2.8). Both indices demonstrated
distinct clustering within communities enriched at the same redox potential with distances
(dissimilarity) becoming greater with each successive redox potential used. These shifts in
community structure were not only dependent on redox potential (Figure 2.8A) but also on the
differences in geochemistry of the two microcosm incubation runs (Figure 2.8B).

Changes in community structure were evaluated by looking at the overall abundance of the
OTUs in each sample along with ordination-constructed heatmaps of the most dominant groups
41
enriched in the cathode-oxidizing communities. The Proteobacteria dominated all electrode
communities, and the second most abundant phylum in all electrode communities was the
Firmicutes. Specifically, the enrichment of Firmicutes was comprised of members of the
Clostridia. While much of the literature has focused on EET mechanisms in the Proteobacteria,
there have been numerous reports of Clostridium strains being electrochemically active (Köpke
et al. 2010; Choi et al. 2012; Choi et al. 2014). Many of these strains are acetogenic bacteria,
capable of using electrons supplied by a cathode to fix carbon dioxide to acetate (Nevin et al.
2011). The mechanisms through which these Clostridium strains perform EET is still unclear and
further investigation into the EET capability of Clostridia is needed as they are consistently
abundant members enriched in electrochemical systems (Tender et al. 2002; Torres et al. 2009;
Dennis et al. 2013; Bretschger et al. 2015). OTUs within the Desulfobacterales were enriched
among all cathode-oxidizing microbial communities. The order was highly abundant across all
electrode communities, comprising 24.55% to 58.64% of the total abundance of the top 50 OTUs
at the order level. In a recent study, the potential mechanisms of how sulfate reducing bacteria
(SRB) receive electrons from their synthrophic anaerobic methanotrophic archaea partners was
investigated (Skennerton et al. 2017). The study identified large multiheme cytochromes in SRB
genomes from members of the Desulfobacterales and Desulfuromonadales that may be involved
in receiving extracellular electrons (Skennerton et al. 2017). Interestingly, the
Desulfuromonadales was another order enriched among the cathode-oxidizing microbial
communities. Members of the Desulfuromonadales are known metal reducing organisms
(Lovley et al. 2004), and some strains also have the ability to accept electrons from electrodes
(Gregory et al. 2004; Rabaey et al. 2007; Rosenbaum et al. 2011). The flexibility to not only
respire insoluble substrates but also oxidize them may confer an advantage in environments
42
where redox gradients can change at any moment (e.g., sediments with burrowing invertebrates).
The Altermonadales is another order containing metal reducing bacteria that can receive
electrons from an electrode (Hsu et al. 2012; Rowe et al. 2017). The -750 mV (vs. Ag/AgCl)
redox potential had the highest amount of electron uptake activity in sediment microcosms with
electrode communities that were the most highly enriched in Altermonadales OTUs. Some of the
electrochemical isolates from our initial study belong to the Altermonodales, indicating that the
enriched OTUs in the electrode communities are likely active players in cathode oxidation.

Metagenome predictions obtained for the different communities showed significant increases in
specific KEGG pathways associated with electrode communities. Motility related pathways were
markedly found in greater abundances in the electrode communities. The ability to be
chemotactic and motile might be critical for some microorganisms to be able to sense and seek
out the electrode as a source of available energy. The dissimilatory metal reducing bacterium,
Shewanella oneidensis MR-1 was shown to have increased swimming speeds around insoluble
substrates (Harris et al. 2010), and inhibition in sensing the presence of insoluble electron
acceptors with the deletion of genes involved in chemotaxis (Harris et al. 2012). The OTUs
contributing the most to the abundance of the motility KOs were members of the
Desulfobacterales and the Desulfuromonadales. Both of these orders were some of the most
highly enriched in the cathode-oxidizing communities (Figure 2.7). Investigating OTU
contributions to the nitrate reductase KEGG ortholog, K02570 not only highlighted groups
(Desulfobacterales and Alteromonadales) that were enriched before when looking at OTU
abundances but also the Campylobacterales, a group that was not observed in prior analyses. By
only looking at the top OTUs, there is often a tendency to overlook the rare biosphere of a
community that can be integral to important processes (Sogin et al. 2006, Huse et al. 2010). The
43
Campylobacterales are an order within the Epsilonproteobacteria typically known as pathogens,
but also contain non-pathogenic members that possess the ability to oxidize reduced sulfur
compounds and are found in sulfide rich sediments (Llorens-Marès et al. 2015; Waite et al.
2017). The enriched OTUs within the Campylobacterales of the -500 and -600 mV (vs.
Ag/AgCl) electrode communities could also be important players in EET processes.

CONCLUSION
Electrochemical enrichment techniques provide a valuable way to study microorganisms capable
of extracellular electron transport. Using this electrochemical approach, we were able to enrich
for cathode-oxidizing microbial communities at seven different redox potentials. The changes
observed in community structure are related to increasing niche availability by providing
cathodic redox potentials (electron donors) with energetics (coupled to electron acceptors)
driving shifts in composition. Various OTUs within the Desulfobacterales, Desulfuromondales,
Rhodobacterales, Altermonadales, and Campylobacterales are candidates for organisms
performing cathode oxidation. Further investigation is necessary to determine which of the
enriched groups are actually involved in EET activity. This study provides an essential starting
point for further understanding and targeted culturing of microorganisms capable of insoluble
substrate oxidation.

44
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50
CHAPTER 3

ELECTROCHEMICAL ENRICHMENT AND ISOLATION OF NOVEL
MICROORGANISMS CAPABLE OF CATHODE OXIDATION FROM MARINE
SEDIMENT

INTRODUCTION

The discovery that microorganisms can uptake electrons supplied by a cathode has significant
ecological and industrial importance. Microorganisms with extracellular electron transport (EET)
ability to receive or donate electrons from insoluble substrates are integral players when it comes
to the biogeochemical cycling of sulfur and iron (Emerson et al. 2010; Holmkvist et al. 2011; Shi
et al. 2016). The capability of microorganisms to use insoluble substrates as electron donors
could help explain why microbial life exists thousands of meters below the seafloor in some of
the most oligotrophic regions of the world (Jørgensen & D’Hondt 2006; D’Hondt et al. 2009;
Hoehler & Jørgensen 2013). Direct measurements of microbial metabolism in the deep dark
ocean far exceed the expected models based on the influx of organic carbon to the system (Burd
et al. 2010). Metabolisms such as lithotrophy that depend less on organic carbon, yet potentially
provide a source of organic carbon may account for some of the missing carbon in current global
budgets (Swan et al. 2011; Hoehler & Jørgensen 2013). The application of electrochemical
techniques to microbiology is enabling us to study EET processes in environmental systems with
electrodes serving as analogs to insoluble substrates (Rowe et al. 2015; Jangir et al. 2016).

The first report of cathode oxidation was in a study by Gregory et al. in 2004 where electrodes
were provided as the sole electron donor coupled to the reduction of nitrate to nitrite and to the
reduction of fumarate to succinate by two Geobacter species. Since then there has been an
increased focus on utilizing microorganisms to catalyze the cathode reactions in microbial fuel
51
cell (MFC) systems (Clauwaert, Van Der Ha, et al. 2007; Chen et al. 2008). Typically the most
effective catalysts for cathodic reactions in MFCs have been materials (e.g. platinum) that are
prohibitive due to economic cost. Microbial mediated catalysis of cathodic reactions is an
alternative to expensive abiotic catalysts, and may provide economic and environmental
advantages when it comes to the generation of useful chemicals through cathodic chemical
synthesis (Rozendal et al. 2009; Foley et al. 2010). The field of microbial electrosynthesis, in
which microorganisms utilize electrons from cathodes to produce organic compounds from CO
2
,
is of particular interest due to the potential of sustainable product generation and synthesis of
biofuels (Rabaey & Rozendal 2010; Tremblay & Zhang 2015). An increased understanding of
what microorganisms are capable of cathode oxidation and how they are performing EET is
necessary to optimize these possible industrial applications.

Most enrichments of cathode-oxidizing microorganisms have originated from applied systems
such as wastewater or MFCs (Clauwaert, Rabaey, et al. 2007; Rabaey et al. 2008; Wrighton et al.
2010). Our laboratory’s pilot study was the first report of an enrichment of cathode-oxidizing
microorganisms from an environmental system (Rowe et al. 2015). Natural environments with
reduced, solid compounds (e.g., FeS, elemental sulfur) have been shown to harbor a high
potential of microorganisms with cathode oxidation capabilities (deCamposRodrigues &
Rosenbaum 2014). Our initial study resulted in the successful isolation of several
electrochemically active microorganisms capable of cathode oxidation from marine sediment
(Rowe et al. 2015). In this work, we further expand the repository of environmental isolates with
cathode oxidation ability. A three-electrode electrochemical enrichment approach utilizing
several redox potentials was combined with traditional culturing techniques to isolate six novel
strains from Catalina Harbor sediment (Catalina Island, CA). Insights into the dynamics between
52
biofilm and planktonic cathode-oxidizing communities were also investigated through molecular
approaches. Electrochemical characterization of the various isolates suggests that these strains
perform EET with distinct and unique mechanisms. The results of this study provide additional
evidence that cathode oxidation is far more prevalent and may be an ability that is more
widespread amongst microorganisms than previously thought.

MATERIALS & METHODS

Enrichment Medium Composition for Secondary Bioreactors and Electrochemical Tests

An artificial seawater (ASW) base served as the medium for all secondary bioreactors and
enrichments. The media was prepared per Rowe et al. (2015). The artificial seawater medium
contained 342 mM NaCl, 14.8 mM MgCl
2
⋅6H
2
O, 0.1 mM CaCl
2
⋅2H
2
O, and 6.7 mM KCl.
Nitrogen, phosphorous, and sulfur were added to the seawater base to yield concentrations of 10
mM ammonium chloride, 1 mM potassium phosphate (pH 7.2), and 1 mM sodium sulfate.
Sodium bicarbonate was added to all secondary bioreactor and anaerobic media at a
concentration of 5 mM to support autotrophic growth.

Figure 3.1. Three-electrode sediment free bioreactors used for secondary enrichment.
A) Diagram of the three-electrode system set-up used for secondary enrichment in the
bioreactors and B) bioreactors incubated in the anaerobic chamber.

B)
A)
53
Secondary Electrochemical Enrichment in Sediment Free Bioreactors

Working indium tin oxide (ITO) electrodes from the sediment microcosm incubation
experiments of Chapter 2 along with electrodes from another microcosm enrichment conducted
(June, 2015) were stored and transferred in anaerobic stoppered bottles filled with artificial
seawater base. These electrodes served as the working electrodes in three-electrode sediment free
bioreactors used for secondary enrichment of cathode-oxidizing microorganisms (Figure 3.1A).
The bioreactors were assembled and autoclaved, and all subsequent set-ups were conducted in an
anaerobic chamber (Coy Laboratory Products, Inc., Grass Lake, MI) (Figure 3.1B). The working
electrodes were poised at the same redox potential they were set at during initial sediment
microcosm incubation; all redox potentials reported here are vs. Ag/AgCl. Ag/AgCl reference
electrodes (1M KCl, BASi, West Lafayette, IN) and platinum counter electrodes were used in the
three-electrode systems. Reference electrodes were sterilized with 70% ethanol and UV prior to
use. Each secondary bioreactor contained 250 mL artificial seawater media (described above).
Applied redox potential was controlled using an eDAQ quad channel potentiostat (eDAQ Inc.,
Colorado Springs, CO). Redox potential and current production were controlled and recorded
with the eCorder eCHART software every 5 minutes (eDAQ Inc., Colorado Springs, CO).

The bioreactors were provided with either sulfate or nitrate to serve as the terminal electron
acceptor (TEA). Geochemical analyses of the sediment microcosms and 16S rRNA community
analysis of initial electrode biofilms provided insight into sulfate and nitrate potentially being
important TEAs coupled to electrode oxidation (Chapter 2). With the addition of each TEA (final
concentration of 200 µM Na
2
SO
4
or NaNO
3
) electron uptake was monitored for changes in
current, as negative current is indicative of electrode oxidation.

54
Media Exchange and Analysis of Flavins, Proteins, Anions, and pH

During the course of secondary bioreactor enrichment, media from the bioreactors was removed
and replaced with fresh sterile artificial seawater base every four weeks. Removed media was
analyzed for flavins, total proteins, anions, and pH. Media was filtered through 0.2 µm Acrodisc
Supor® membrane syringe filters (Pall Corporation, Port Washington, NY) and filtrate was
stored at -80°C until analysis for flavins and anions. 2 mL aliquots of media were centrifuged at
13,000 rpm for 5 min to concentrate cells for total protein analysis. Pellets were stored at -20°C
until analysis. Flavin concentrations were measured as described by Woodcock et al. (1982). The
column utilized was a 3.0 mm x 15 mm Zorbax StableBond C18 solvent saver column with a 1.8
µm particle size (Agilent Technologies, Santa Clara, CA). A fluorescence detector (Agilent
Technologies, Santa Clara, CA) was used with an excitation wavelength of 450 nm and an
emission wavelength of 520 nm. To measure total proteins, pellets were hydrolyzed in 1 M
NaOH at 50°C for 30 min with vigorous mixing by vortex at 10 min intervals. Soluble protein in
the extracts was determined spectrophotometrically using Folin phenol reagent (Lowry et al.
1951) with bovine serum albumin (BSA) as a standard. Anion analysis was performed using a
Metrohm Ion Chromatograph (Metrohm, Riverview, FL) with an anions seawater separation
column (Metrosep A SUPP 250, Metrohm). The ion chromatograph was run as per manufacturer
protocol with 3.2 mM sodium bicarbonate, 1 mM sodium carbonate running buffer containing
2% acetonitrile at a flow rate of 0.7 mL/min.

DNA Extraction and 16S rRNA Community Analysis

For DNA extraction, biofilm from the electrodes was scraped off and electrodes were treated
with sonication in sterile artificial seawater base to remove as much biomass as possible. In
addition, electrodes were treated with 200 µl of lysozyme and 200 µl proteinase-K to remove any
55
biofilm remaining on the ITO coated glass. Planktonic cells were collected by centrifuging 50
mL of media from the bioreactors at 10,000 rpm for 10 min at 4°C. Resultant cell pellets,
electrode biomass and enzymatic treatment were directly added to bead beating tubes from a MO
BIO PowerSoil DNA Isolation Kit (Qiagen, Carlsbad, CA). DNA extraction was performed
using the MO BIO Powersoil DNA Isolation Kit according to manufacturer’s protocols.
Extracted DNA quality and quantity was determined using a Nanodrop (Thermo Fisher
Scientific, Wilmington, DE).

Illumina MiSeq paired-end sequencing was performed on the V4 hypervariable region of the 16S
rRNA gene of extracted DNA by MR-DNA (Molecular Research LP, Shallowater, TX). Briefly,
ribosomal sequences were amplified with a 30-cycle PCR reaction with a 53°C annealing temp
using barcoded 515F primer paired with 806R primer. Amplicons were pooled and purified with
calibrated Agencourt Ampure XP beads (Agencourt Bioscience Corporation, Beverly, MA).
Purified PCR product was used to prepare DNA libraries by following the Illumina TruSeq DNA
library preparation protocol and sequenced on an Illumina MiSeq following manufacturer’s
guidelines (Illumina Inc., San Diego, CA). Average sequence lengths of 250 bp and > 70,000
sequences were obtained for all samples.

QIIME (Quantitative Insights Into Microbial Ecology) bioinformatics pipeline (Caporaso et al.
2010) was utilized for quality filtering (Bokulich et al. 2013), chimera checks (Edgar et al.
2011), and barcode removal. De novo OTU (operational taxonomic unit) picking was used for
taxonomy assignment (based on 97% similarity), sequence alignment (Caporaso et al. 2010), and
phylogenetic analysis using the Greengenes database (Werner et al. 2012). The R package,
Phyloseq (McMurdie & Holmes 2013) was implemented for downstream analysis of 16S rRNA
56
sequencing data including non-metric multidimensional scaling (NMDS). NMDS plots were
based on normalization of reads of community samples to account for differences in sequencing
depth. Permutational multivariate analysis of variance (PERMANOVA) was used to evaluate the
contribution of different variables to bioreactor microbial community variation with the Adonis
function (Anderson 2001).

Genomic Predictions Using 16S rRNA Sequences
Functional composition of the microbial communities from each bioreactor sample was predicted
using the Phylogenetic Investigation of Communities by Reconstruction of Unobserved States
(PICRUSt v. 1.1.2) bioinformatics software package (Langille et al. 2013). Gene family
estimates were based on an OTU table generated from 16S rRNA sequences processed by
closed-reference OTU picking against the Greengenes database through QIIME (Caporaso et al.
2010). The OTU table was rarefied to the same number of sequences per sample (n = 63,992).
Functional predictions were based on KEGG (Kyoto Encyclopedia of Genes and Genomes)
orthologs (KO) at two hierarchical levels. Statistically significant differences between KO
abundances in different sample groups (redox potentials or biofilm/planktonic) were determined
using nonparametric Kruskal-Wallis tests.

Tertiary Enrichment Using Insoluble Electron Donor Media and Isolation

Various media containing insoluble electron donors were utilized to further enrich for cathode-
oxidizing microorganisms. Iron was provided in the form of elemental iron granules (1-2 mm,
99.98% metals basis, Alfa Aesar, Haverhill, MA). The iron granules served as the electron donor
source in anaerobic seawater media supplemented with either nitrate or sulfate as the TEA.
Elemental sulfur was prepared as described in Moser and Nealson (1996) and added to artificial
57
seawater media to make agar plates with high (30 mM) and low sulfur (3 mM) concentrations
(1.5% agar). The elemental sulfur agar plates were made with either nitrate or Fe(III)-NTA as
TEAs. Difco marine broth (Difco, Lawrence, KS) was also utilized as an enriched media to
screen for isolates that could also grow heterotrophically.

Chronoamperometry and Cyclic Voltammetry of Isolates

All electrochemical tests were conducted in three-electrode electrochemical cells (15 mL
volume). ITO plated glass or carbon felt was used as the working electrode, with a platinum wire
counter electrode, and a 1M KCl Ag/AgCl reference electrode. Chronoamperometry was
performed on isolates using the eDAQ quad channel potentiostat and eCORDER eCHART
software as described above. All isolates were grown up overnight in enriched marine broth at
30°C (Difco, Lawrence, KS). After 24 hr, 20 mL of the overnight culture was pelleted at 13,000
rpm for 10 min and washed and resuspended in ASW base. The resuspended biomass was added
directly to electrochemical cells sparged with filtered air or nitrogen at a rate of 20 mL/min. If
nitrate was used as the TEA, sodium nitrate was provided at a concentration of 200 µM in batch
feeds. The electrochemical cells were either poised at +100 mV (vs. Ag/AgCl) to initiate biofilm
formation or directly exposed to cathode-oxidizing conditions at -400 to -600 mV (vs. Ag/AgCl).
Air flow was stopped at certain periods to see if electrode oxidation could be coupled to oxygen
reduction. Futhermore, cyanide injections were done to look for inhibition in electrode oxidation.
Media from the electrochemical cells was sampled during the course of chronoamperometry runs
to also assay for flavins as described for secondary bioreactor media analysis. Cyclic
voltammetry (CV) tests were conducted on the direct biofilms of isolates grown in the
electrochemical cells or planktonic cells removed from the electrochemical cells to characterize
the electrochemical activity of each fraction. In addition, filtrate from the media was also
58
examined through cyclic voltammetry. CVs were performed with a Gamry Reference 600
potentiostat and Gamry Framework software (Gamry Instruments, Warminster, PA). Each CV
analysis was conducted at a scan rate of 5 mV/sec over a range of -800 to 800 mV (vs. Ag/AgCl)
Analysis on CV curves was performed with Gamry Echem Analyst (Gamry, Warminster, PA)
including calculation of mid-point potentials.

Scanning Electron Microscopy (SEM) Sample Preparation

Planktonic cells were immediately filtered through sterile 0.2 µm Supor® 200 filters (Pall
Corporation, Port Washington, NY). Filters and electrode samples were fixed in distillation
purified electron microscopy grade 2.5% glutaraldehyde (Electron Microscopy Sciences,
Hatfield, PA). Samples were then processed through an ethanol dehydration series (30, 50, 70,
80, 90, 95, 100% v:v ethanol, three 10 min intervals for each concentration) and critical point
drying (Autosamdri 815 critical point drier, Tousimis Inc., Rockville, MD). Samples were
mounted on aluminum stubs and coated with Au (Sputter Coater 108, Cressington Scientific,
Watford, UK). Images were captured at 5 keV using a JEOL JSM 7001F low vacuum field
emission scanning electron microscope (JEOL USA, Inc., Pleasanton, CA).

Phylogenetic Analysis of Isolates

For taxonomic classification of isolates, stocks were streaked on enriched seawater agar plates
(Difco, Lawrence, KS). PCR was performed directly on colonies or DNA extracted from pellets
of overnight liquid cultures to amplify the 16S rRNA gene with the primers 27F and 1492R.
Purified PCR product (DNA Clean & Concentrator™-5, Zymo Research, Irvine, CA) was
sequenced (Genewiz, South Planfield, NJ) with both primers. Sequences were quality checked
using 4Peaks (Nucleobytes, Netherlands) and aligned against the SILVA database using the
59
SINA aligner (v 1.2.11) (Pruesse et al. 2012; Quast et al. 2013). Sequences from isolates from
our laboratory’s previous work (Rowe et al. 2015) and nearest related organisms were also
included in alignments. Maximum likelihood phylogenetic trees were built using the RAxML
program v.8 (Stamatakis 2014) with the ARB software package interface (Ludwig et al. 2004).

RESULTS
Secondary Bioreactor Electrochemical Enrichments: A total of 5 secondary bioreactors were
constructed and run with different redox potentials and electron acceptors as described in Table
3.1 for further enrichment of cathode-oxidizing microorganisms. All secondary bioreactors
exhibited electron uptake by the cells (increases in negative current) when supplemented with a
terminal electron acceptor (TEA). Figure 3.2 shows typical responses in current in the secondary
bioreactors when fed with a TEA. There were no changes in current when bioreactors were not
supplied with a TEA, suggesting that cathode oxidation only occurred when microorganisms had
an available TEA to couple with electrode oxidation. The performance of the secondary
bioreactors evaluated as average current generation varied from -1.12 to -29.5 µA. Despite
differences in current generation during initial microcosm enrichment (Chapter 2, Figure 2.4)
most secondary bioreactors produced current in a similar range.

Table 3.1. Summary of secondary bioreactor general run information

*Denotes bioreactor stopped due to working electrode short-circuiting
60

Figure 3.2. Chronoamperometry profiles of secondary bioreactors. A) -650 mV (vs.
Ag/AgCl) bioreactor fed with sulfate as the TEA and B) -400 mV (vs. Ag/AgCl) (orange)
and -500 mV (vs. Ag/AgCl) (pink) bioreactor fed with nitrate as the primary TEA. Green
arrows indicate sulfate feeds, red arrows indicate nitrate feeds, and the black dashed arrow
indicates an artificial seawater base feed. Sulfate and nitrate additions correspond to a final
concentration of 200 µM.
61
Table 3.2. Average pH, proteins, flavins, nitrite and sulfate concentrations in media from
secondary bioreactor media exchanges

*Denotes only one sample was obtained for measurement
#
Denotes only one sample exhibited detectable concentrations
ND indicates not detected

Media exchanges were performed during the course of the run for the secondary bioreactors with
the longest run time (-550, -400, and -500 mV vs. Ag/AgCl). Measurements of pH, proteins,
flavins, and anions were taken of media removed (Table 3.2). A decrease in pH was observed for
all secondary bioreactors as the ASW media used was initially made to reflect the pH of seawater
collected from Catalina Harbor at a pH of 8.2. Protein concentrations in the media had high
variability during the run, ranging from 10.55 to 362.301 µg/ml. High protein concentrations
generally occurred at points where negative current generation was the most robust (data not
shown). Even though each media exchange resulted in the removal of the planktonic cells from
the bioreactors, protein concentrations indicate that the planktonic communities were able to
recover and are most likely seeded from the electrode biofilm communities. Flavins in the form
of flavin mononucleotide (FMN) and riboflavin (RF) were measured for all media exchanges.
FMN was consistently detected across all bioreactors while RF was only detected once in the
-500 mV (vs. Ag/AgCl) bioreactor (Table 3.2). These flavins have been implicated to play a role
in EET, serving as electron shuttles between microorganisms and insoluble substrates (Marsili et
al. 2008; von Canstein et al. 2008; Okamoto et al. 2013). It is becoming clear that in some cases
these flavins interact directly with multiheme cytochrome proteins to facilitate EET (Okamoto et
al. 2014; Edwards et al. 2015). Recently, flavins have been detected and measured in marine
62
sediments, with FMN being found in nanomolar concentrations (0.4 – 1.2 nM) (Monteverde et
al. 2016). The constant presence of FMN found in secondary bioreactors suggests that this flavin
might be important for EET activity in the bioreactor communities.

Figure 3.3. SEM images of the planktonic cells from two of the secondary enrichment
bioreactors. A) & B) planktonic cells from -400 mV (vs. Ag/AgCl) secondary bioreactor, C)
& D) planktonic cells from the -500 mV (vs. Ag/AgCl) secondary bioreactor.

Concentrations of nitrite, nitrate, and sulfate were measured as sulfate and nitrate served as TEA
for the secondary bioreactors. The -550 mV (vs. Ag/AgCl) bioreactor was supplemented with
sulfate as the TEA, while the -400 and -500 mV (vs. Ag/AgCl) bioreactors were supplied with
nitrate as the TEA. During the course of a month between media exchanges, approximately 1
A) -400 mV Planktonic

B) -400 mV Planktonic

C) -500 mV Planktonic

D) -500 mV Planktonic

63
mM of the corresponding TEA was added to each bioreactor. Sulfate was found in trace amounts
in the -550 mV (vs. Ag/AgCl) bioreactor. Additionally, ASW media used for the bioreactors had
a baseline concentration of 1 mM sodium sulfate. The low levels of sulfate measured (0.04 mM)
indicate the bioreactor communities were actively using sulfate as a TEA. The -400 and -500 mV
(vs. Ag/AgCl) bioreactors also exhibited low levels of sulfate (0.17 – 0.19 mM) and nitrite in the
0.49 mM range. Although sulfate did not serve at the primary TEA for these bioreactors, there
was a low level response in negative current when sulfate was added (Figure 3.2B). Other
members of the bioreactor communities not actively engaging in cathode oxidation could also
potentially use sulfate. Nitrite measured in the bioreactor media could be a byproduct of nitrate
reduction. Other byproducts of denitrification (e.g. NO, N
2
O) were not measured and could
account for nitrate utilization. Cells present in the bioreactor media were collected and fixed for
imaging with SEM (Figure 3.3). Diverse morphologies and cell sizes were observed in the -400
and -500 mV (vs. Ag/AgCl) planktonic communities. In some cases, thin extracellular filaments
resembling nanowire appendages connecting cells to one another were present (Figure 3.3D).
Nanowire appendages have been reported to be a mode of EET that allow microbial cells to
perform electron transfer at a distance (Reguera et al. 2005; Gorby et al. 2006).

Secondary Bioreactor Communities: Microbial community analyses were performed on final
electrode biofilm communities and planktonic communities with amplicon next generation
sequencing of the 16S rRNA gene. Overall richness (estimated by observed and Chao1 indices)
and diversity (estimated by Shannon and Simpson indices) was higher in electrode biofilm
communities compared to planktonic communities across all redox potentials (Figure 3.4). For
the secondary bioreactors (-550, -400, and -500 mV vs. Ag/AgCl) where multiple planktonic
samples were taken, richness and diversity also increased with time. The redox potentials that
64
had shorter run times (-650 and -750 mV vs. Ag/AgCl) displayed lower richness and diversity
measures compared to the longer running redox potentials (-550, -400, and -500 mV vs.
Ag/AgCl).

Figure 3.4. Species richness (observed and Chao1) and diversity (Shannon and Simpson)
measures of the bioreactor communities from A) -550, -650, and -750 mV (vs. Ag/AgCl)
secondary bioreactors and B) -400 and -500 mV (vs. Ag/AgCl) secondary bioreactors.

65

Figure 3.5. Relative abundances for each bioreactor community sample at the class (top
100 OTUs) and order (top 50 OTUs) level. The -550, -650, and -750 mV (vs. Ag/AgCl)
secondary bioreactor communities are shown in A) at the class level and in B) at the order level.
The -400 and -500 mV (vs. Ag/AgCl) bioreactor communities are shown in C) at the class level
and D) at the order level.

66
Shifts in community structure between redox potentials and mode of growth (planktonic vs.
biofilm) were evaluated in terms of OTU abundance (Figure 3.5). For the secondary bioreactors
where sulfate served as the TEA (-550, -650, and -750 mV vs. Ag/AgCl), there was an overall
increase in Deltaproteobacteria in the electrode biofilm communities and an increase in
Gammaproteobacteria in the planktonic communities across all redox potentials. The most
dominant members in the communities also changed with redox potential. At the class level, the
-550 mV (vs. Ag/AgCl) bioreactor had an enrichment of Mollicutes (4.16%-15.29%) that is not
observed with the other redox potentials. The -750 mV (vs. Ag/AgCl) redox potential had the
highest overall enrichment of Epsilonproteobacteria (52.38%-58.98%) while the -650 mV (vs.
Ag/AgCl) redox potential had greatest enrichment of Deltaproteobacteria (45.25%-72.65%). The
secondary bioreactor communities where nitrate served as the TEA, -400 and -500 mV (vs.
Ag/AgCl), are much more dominated by the Gammaproteobacteria compared to the sulfate
secondary bioreactors. The Gammaproteobacteria comprised 77.29%-77.64% of the -400 mV
(vs. Ag/AgCl) electrode biofilm communities. In comparison, the highest abundance of
Gammaproteobacteria in the sulfate secondary bioreactor communities was 36.29% in the -650
mV (vs. Ag/AgCl) planktonic community. We were unable to obtain an electrode biofilm sample
for the -500 mV (vs. Ag/AgCl) bioreactor, because the electrode from that reactor experienced
short-circuiting. Changes in community structure between the -400 mV (vs. Ag/AgCl) electrode
and biofilm communities differ from the trends seen with the sulfate secondary bioreactors.
Electrode communities exhibited a notable increase in the Alphaproteobacteria and to a lesser
extent the Gammaprotebacteria. The planktonic communities had a higher presence of the
Epsilonproteobaceria and Deltaproteobacteria as compared to electrode assemblages.

67
Abundances at the order level for each set of secondary bioreactors give deeper insight into what
specific microbial groups were enriched (Figure 3.5B,D). The Campylobacterales and
Alteromonadales were consistently found across all redox potentials and communities. The
Campylobacterales are an order within the Epsilonproteobacteria. Epsilonproteobacteria are
commonly found in deep-sea hydrothermal vent and subsurface systems and are noted for their
chemolithotrophic metabolisms (López-García et al. 2003; Takai et al. 2005; Campbell et al.
2006; Yamamoto & Takai 2011). Epsilonproteobacteria have been found to colonize the
surfaces of carbon steel and stainless steel substrates submerged in coastal seawaters (Jones et al.
2007; Bermont-Bouis et al. 2007). Several Campylobacterales OTUs were reported to be found
on corroded carbon steel incubated in Pacific Ocean seawater (Dang et al. 2011). The sulfate
secondary bioreactor communities, which exhibited increases in the Deltaproteobacteria, had
enrichments of the Desulfovibrionales (8.24%-67.71%) and Desulfobacterales (2.32%-19.50%).
On the other hand, the nitrate secondary bioreactor communities were highly enriched in the
Chromatiales (11.36%-38.92%). Chromatiales OTUs have been identified to be dominant
members of denitrifying cathodes from fuel cell systems (Wrighton et al. 2010). Recently, a
member of the Chromatiales was shown to be an important constituent of a biocathode enriched
from seawater with studies suggesting it performs CO
2
fixation driven by cathode oxidation
(Leary et al. 2015; Wang et al. 2015; Yates et al. 2016).

Differences in overall community structure were quantified with hierarchical clustering using
Bray-Curtis dissimilarity matrices (Figure 3.6). The NMDS plot demonstrates differences in
microbial community structure with samples that are spatially closer together showing greater
similarity. Overall, bioreactor communities from the same redox potential showed the closest
similarity to each other. It is worth noting, that although the communities group together based
68
on redox potential, the electrode biofilm and planktonic communities generally do not overlap.
The exception to the aforementioned trend is with the -400 mV (vs. Ag/AgCl) redox potential
communities. The -400 mV (vs. Ag/AgCl) electrode biofilm communities share great similarity
to the later -400 and -500 mV (vs. AgCl) planktonic communities. A drastic shift in the
planktonic communities of the -400 and -500 mV (vs. Ag/AgCl) secondary bioreactors took
place after the 3/4/16 sampling date. The more negative redox potentials, -650 and -750 mV (vs.
Ag/AgCl) also spatially group together while the other redox potentials (-400, -500, and -550
mV vs. Ag/AgCl) are found to cluster closer together as well.

Figure 3.6. Bray-Curtis based non-metric multidimensional scaling (NMDS) plot of the
secondary bioreactor community samples. Circles represent electrode biofilm samples and
triangles signify planktonic samples.
69
Adonis tests were performed on weighted Unifrac values to determine the significance of
variation observed in the microbial communities explained by growth mode (planktonic vs.
biofilm), redox potential, and terminal electron acceptor. The mode of growth did not exhibit a
significant impact on the variation between the microbial communities. Redox potential
contributed the most to variation (R = 0.71, p = 0.001), followed by the terminal electron
acceptor used (sulfate or nitrate) (R = 0.44, p = 0.001).

Genomic Potential of Bioreactor Communities: Genome predictions of the secondary bioreactor
microbial communities were estimated with PICRUSt. PICRUSt determines the phylogenetic
distance between predicted metagenomes and reference genomes as a way to assess the accuracy
of the metagenome predictions by calculating nearest sequenced taxon index (NSTI) values
(Langille et al. 2013). Smaller NSTI values reflect a higher accuracy of prediction. NSTI values
for secondary bioreactor communities varied from 0.062-0.15. All of the NSTI values obtained
fall within the range reported for accurate metagenome predictions with NSTI values ≤ 0.17

Table 3.3. Relative abundances of predicted KEGG orthologs assigned to KEGG pathways
in bioreactor community samples found to have significant differences in abundance.
Biofilm communities are designated Elec, and planktonic communities are designated Plank.

*Denotes a significant difference (p < 0.05) in the abundance of a KEGG pathway grouping
associated with biofilm and planktonic communities by Kruskal-Wallis tests. All other KEGG
pathways have a significant difference (p < 0.05) in abundance based on redox potential
determined by Kruskal-Wallis tests.
70
(Langille et al. 2013). The relative abundance of relevant KEGG pathways associated with
metabolism and cellular processes/signaling are reported in Table 3.3. All of the pathways listed
in Table 3.3 were found to have a significant difference (p < 0.05) in abundance between
bioreactor communities either based on redox potential or association with the biofilm or
planktonic state. Methane metabolism was found in higher abundances in the electrode
communities from the sulfate secondary bioreactors, and was especially high in the -650 mV (vs.
Ag/AgCl) redox potential communities. The -650 mV (vs. Ag/AgCl) electrode community had
an enrichment of Methanomicrobia (Figure 3.5). There have been reports of methane production
by biocathode communities enriched in Methanobacterium spp. provided with only the cathode
as an electron donor and CO
2
as the sole carbon source (Cheng et al. 2009; Marshall et al. 2012).
All other pathways, including nitrogen and sulfur metabolism, were found in differential
abundance based on redox potential. The -750 mV (vs. Ag/AgCl) redox potential communities
had particularly high abundances in nitrogen metabolic pathways relative to other redox
potentials. Genes related to electron transfer carriers were the most abundant in the -550 mV (vs.
Ag/AgCl) redox potential communities and the -650 mV (vs. Ag/AgCl) electrode community.
Metabolism of cofactors and vitamins pathways were increased in the -750 mV (vs. Ag/AgCl)
planktonic community, and oxidative phosphorylation genes were found in the greatest
abundance in both -750 mV (vs. Ag/AgCl) redox potential communities.

As both sulfur and nitrogen metabolism pathways were found to have significant differences in
abundance, specific KEGG orthologs (KOs) related to adenylylsulfate reductase and nitrate
reductase were investigated in terms of OTU contribution to KOs (Figure 3.7). These specific
orthologs (K00394, adenylylsulfate reductase and K00370, nitrate reductase) were chosen due to

71

Figure 3.7. Abundance of KEGG Orthologs related to A) Adenylylsulfate reductase
(K00394, adenylylsulfate reductase, subunit A) and B) nitrate reductase (K00370, nitrate
reductase 1, alpha subunit) in each microbial community contributed by OTUs at the order
level.
72
the fact that sulfate and nitrate both served as the primary TEAs in the secondary bioreactors.
Adenylylsulfate reductase performs the second step in dissimilatory sulfate reduction, reducing
adenosine 5’-phosphosulfate to sulfite. The orders accounting for most of the presence of
K00394 in the bioreactor communites were the Desulfovibrionales for the sulfate secondary
bioreactors and the Desulfobacterales and unknown orders for the nitrate secondary bioreactors
(Figure 3.7A). Interestingly, only the -650 mV (vs. Ag/AgCl) bioreactor communities had a
significantly higher abundance of K00394 relative to the nitrate secondary bioreactors. In the
case of nitrate reductase, the -400 and -500 mV (vs. Ag/AgCl) bioreactor communities had
markedly higher abundances compared to the sulfate secondary bioreactor communities for the
most part. The -550 mV (vs. Ag/AgCl) secondary bioreactor, was the exception exhibiting an
increased level of K00370. The orders contributing to the presence of nitrate reductase differed
based on secondary bioreactor TEA. For the nitrate fed secondary bioreactors, -400 and -500
mV (vs. Ag/AgCl), the Chromatiales and Alteromonadales were the primary contributors to the
occurrence of nitrate reductase. With the sulfate fed secondary bioreactors (-550, -650, and -750
mV vs. Ag/AgCl), the Desulfobacterales and Altermonadales were the main drivers for the
abundance of K00370.

Tertiary Enrichment Cultures and Isolation of Cathode Oxidizers: Further enrichment for pure
isolates was approached by seeding tertiary enrichment cultures with samples from the secondary
enrichment bioreactors. Media were designed with iron granules as the electron donor source and
sulfate or nitrate as the sole electron acceptor (Figure 3.8C). After several passages, the
microbial community structure of the Fe
0
/SO
4
tertiary enrichments was evaluated with 16S
rRNA sequencing (Figure 3.8). The tertiary enrichment culture seeded from the -550 mV (vs.
AgCl) redox potential secondary bioreactor harbored a microbial community distinctly different
73
from the -650 and -750 mV (vs. Ag/AgCl)-seeded cultures. The -550 mV (vs. Ag/AgCl) tertiary
enrichment culture was almost exclusively comprised of Gammaproteobacteria belonging to the
order Alteromonadales. The -650 and -750 mV (vs. Ag/AgCl) tertiary enrichment cultures were
dominated by three orders of Deltaproteobacteria: Syntrophobacterales, Desulfuromonadales,

Figure 3.8. Relative abundance at the A) class and B) order levels (top 150 OTUs) of
Fe
0
/SO
4
tertiary enrichment cultures, and C) photograph of Fe
0
/SO
4
tertiary enrichment
culture serum vials. Enrichment cultures were initially inoculated with media from the -550,
-650, and -750 mV (vs. Ag/AgCl) secondary bioreactors.

C
74

Figure 3.9. Maximum likelihood phylogenetic tree constructed for all isolates. The 16S
rRNA sequences from isolates obtained from the pilot study are shown in teal (Rowe et al.
2015). The 16S rRNA sequences from strains isolated from this study are shown in red.

and Desulfovibrionales. Clostridia were the second most abundant group in the -650 and -750
mV (vs. Ag/AgCl) enrichment cultures. Isolates from the Fe
0
/SO
4
and Fe
0
/NO
3
tertiary
enrichment cultures were obtained through dilution to extinction in FeCl
2
agar shake tubes.
Isolates utilizing sulfur as an electron donor source were plated on elemental sulfur plates
supplemented with nitrate as an electron acceptor. The plates were initially streaked with
samples from the secondary enrichment bioreactors, and colonies that produced clearing of the
elemental sulfur were chosen for further plating. The iron-oxidizing isolates obtained fell within
the Deltaproteobacteria (Desulfovibrio genus), the Clostridia (Vallitalea genus), and the
Gammaproteobacteria (Marinobacter genus) (Figure 3.9). The sulfur-oxidizing isolates attained
are from the Epsilonproteobacteria (Arcobacter genus) and Gammaproteobacteria

Clostridia
Gammaproteobacteria
Alphaproteobacteria
Deltaproteobacteria
Epsilonproteobacteria
ARB_E4E3FD7, KC876639.1.1480 Vallitalea pronyensis
ARB_1741EE3F, KC668884.1.1491 Vallitalea; uncultured bacterium
ARB_7E903404, KC668853.1.1491 Vallitalea; uncultured bacterium
ARB_AB292847, KC668851.1.1491 Vallitalea; uncultured bacterium
ARB_77457869, AJ866931.1.1467 Desulfovibrio sp. NB21
ARB_7587F4D5, AJ866945.1.1423 Desulfovibrio sp. NC301
ARB_862527D1, GU194165.1.1407 Desulfovibrio sp. Col 20
ARB_9CA9E356, Campylobacter jejuni
ARB_EFB5AE97, AIPM01000049.478.1994 Campylobacter jejuni subsp.
ARB_D8A9228C, KT952702.1.1486 Arcobacter; uncultured bacterium
ARB_D13629C1, AJ271653.1.1508 Arcobacter sp. B4b1
ARB_EE8950A7, AB013832.1.1477 Arcobacter; uncultured bacterium
ARB_64B55766, KC527432.1.1217 Arcobacter; uncultured bacterium
ARB_3BCB47E9, JN596128.1.1355 Arcobacter; bacterium A2(2011b)
ARB_7E02DD4D, HQ326308.1.1430 Thioclava; uncultured bacterium
ARB_7E02DD4D, KF995170.1.1430 Thioclava; uncultured bacterium
ARB_CE327C08, KF146553.1.1430 Thioclava; uncultured bacterium
ARB_CE327C08, HQ326264.1.1430 Thioclava; uncultured bacterium
ARB_CE327C08, KF146549.1.1430 Thioclava; uncultured bacterium
ARB_777A52D, AY505529.1.1519 Idiomarina loihiensis
ARB_20C8FAEA, EF198113.1.1504 Idiomarina sp. BSw10042
ARB_D9ED8307, DQ642843.1.1511 Vibrio sp. 40
ARB_A82DFF19, KC433749.1.1412 Vibrio harveyi
ARB_E604B4CB, JN188406.1.1459 Vibrio alginolyticus
ARB_E6E2DA0B, KT989853.1.1474 Vibrio neocaledonicus
ARB_5D016ADB, KP016698.1.1452 Vibrio; uncultured Vibrio sp.
ARB_BDC389A3, GU397400.1.1440 Halomonas alimentaria
ARB_468C86B7, AF211860.1.1493 Halomonas alimentaria
ARB_14B4B395, KM823709.1.1530 Halomonas; uncultured bacterium
ARB_43385578, AY505525.1.1511 Halomonas salina
ARB_88E55135, JQ799076.1.1522 Halomonas boliviensis
ARB_23A96DDD, GU212642.1.1518 Halomonas sp. H184B37
ARB_23A96DDD, AJ564880.1.1518 Halomonas alkaliantarctica
ARB_B0E1BD17, JN160749.1.1485 Marinobacter hydrocarbonoclasticus
ARB_9C814882, JN160746.1.1498 Marinobacter hydrocarbonoclasticus
ARB_B16AB47D, HQ696525.1.1456 Marinobacter; actinobacterium YH66
ARB_C6B963AC, HM573378.1.1460 Marinobacter sp. EB297
ARB_42B4BBD0, JN160750.1.1501 Marinobacter hydrocarbonoclasticus
ARB_EF557937, KJ188006.1.1535 Marinobacter sp. L21−PYE−C22
ARB_53609956, DQ486512.1.1445 Marinobacter sp. DG1305
ARB_78B4EF96, JQ032267.1.1459 Marinobacter; uncultured bacterium
ARB_E1EE53AF, JQ032348.1.1456 Marinobacter; uncultured bacterium
ARB_9CE5C8F7, JN874290.1.1484 Marinobacter; uncultured bacterium
ARB_7E65FB84, JN874017.1.1483 Marinobacter; uncultured bacterium
ARB_24922612, AB617558.1.1425 Marinobacter flavimaris
ARB_1979AA1F, JN874030.1.1483 Marinobacter; uncultured bacterium
ARB_A7D41547, HM142822.1.1501 Pseudomonas; proteobacterium WJQ N
ARB_EFFD0B23, JVYC01000169.3709.5225 Pseudomonas mendocina
ARB_126424CB, KF906576.1.1529 Pseudomonas; uncultured bacterium
ARB_B29FD443, AOBS01000001.3672.5210 Pseudomonas stutzeri NF13
ARB_E35A09F8, JYHV01000034.3658.5203 Pseudomonas stutzeri
ARB_E35A09F8, JYHV01000021.3658.5203 Pseudomonas stutzeri
0.10
ARB_B6B9DC3F, FJ716997.1.1536 Desulfovibrio; uncultured bacterium
ARB_8FF5BFBD, JQGR01001230.195.1701 Vallitalea; bioreactor metagenome
ARB_D743DDD9, GU136555.1.1502 Desulfovibrio;delta proteobacterium
ARB_AE6288A, −750 mV Isolate 2
ARB_8C35772B, −550 mV Isolate 2
ARB_A050CF23, −750 mV Isolate 1
ARB_56347A70, Thalassospira sp. 1
ARB_7C136D8B, Thioclava sp. 1
ARB_548F0E46, Idiomarina sp. 1
ARB_2F9F7334, −550 mV Isolate 1
ARB_A150AA38, Halomonas sp. 2
ARB_79D9FB5B, Halomonas sp. 1
ARB_72415B5B, −500 mV Isolate 1
ARB_28D3EFE, Marinobacter sp. 1
ARB_71F0D6C1, −650 mV Isolate 1
ARB_5FC17970, Pseudomonas sp. 3
ARB_E29C7588, Pseudomonas sp. 2
ARB_BFC7BB72, Pseudomonas sp. 1
ARB_CCF00049, AGQZ01000062.2170.3681 Halomonas boliviensis LC1
Teal: Isolates from -400 mV
Red: Isolates from -500 to -750 mV
75

Figure 3.10. Representative chronoamperometry to test for electrochemical activity of
isolates. A) -550 mV Isolate 1, Vibrio sp. and B) -650 mV Isolate 1, Marinobacter sp.. -550
mV Isolate 1 was initially exposed to anodic conditions before switching to cathodic conditions.
The applied potential for the -650 mV Isolate 1 was changed from -400 to -600 mV (vs.
Ag/AgCl) at the 40 hr time point. Arrows indicate switches in conditions. Blue arrows signify air
and cathodic conditions, orange arrow indicates purging with nitrogen, red arrows indicate
cyanide injections, and black dashed arrow indicates change in applied redox potential.

76
(Marinobacter and Vibrio genera) (Figure 3.9). The isolates were also plated on enriched
seawater agar plates to test for heterotrophic growth. All isolated Gammaproteobacteria strains
were capable of growth in enriched marine media, and these strains were also capable of utilizing
both nitrate and oxygen as electron acceptors. The Deltaproteobacteria strain was capable of
growth with both sulfate and nitrate, and the Epsilonproteobacteria strain demonstrated growth
with both nitrate and oxygen. The Clostridia strain was only capable of growth with nitrate. Our
initial study yielded isolates within the Alphaproteobacteria and Gammaproteobacteria (shown
in teal in Figure 3.9) (Rowe et al. 2015). With this work, we have expanded the diversity of
isolates to the Deltaproteobacteria, Epsilonproteobacteria, and Clostridia (shown in red in
Figure 3.9).

Electrochemical activity of isolates was confirmed with chronoamperometry tests. Isolates
demonstrated cathode oxidation during chronoamperometry tests with oxygen or nitrate as a
terminal electron acceptor (examples in Figure 3.10). In some cases, a slightly positive anodic
potential (100 mV vs. Ag/AgCl) was initially applied to get strains to interact with the electrode
before cathodic conditions were applied (Figure 3.10A). Strains were confirmed to perform
electron uptake above the level of controls (sterile ASW media or killed controls). In addition,
for tests where oxygen was used as a TEA, electrochemical cells were purged with nitrogen for
periods of time to eliminate oxygen present. When no oxygen was provided, negative current
ceased and dropped down to baseline levels (Figure 3.10B). Cyanide was also used to inhibit
cytochrome c oxidase, preventing the transport of electrons from cytochrome c to oxygen (Figure
3.10). Negative current was no longer produced after cyanide was introduced to electrochemical
cells. These results suggest a relationship between cathode oxidation and oxygen reduction.

77

Figure 3.11. Plots of cyclic voltammetry tests conducted under non-turnover conditions of
different fractions for isolates, A) -550 mV Isolate 1, Vibrio sp. and B) -650 mV Isolate 1,
Marinobacter sp.. Separate cyclic voltammetry curves were obtained for planktonic and biofilm
fractions. With both strains, the biofilm associated cells show contribution to electrochemical
activity.
78
Table 3.4. Calculated midpoint potentials of isolates determined by cyclic voltammetry
tests in either planktonic or biofilm associated biomass.

*Denotes a weak but quantifiable signal

Electrochemical Characterization of Isolates and Potential Mechanisms of EET:
Investigation into the mode and potential mechanisms of EET conducted by the isolates was
done with electrochemical and imaging techniques. EET is known to occur through direct
contact between the microbial cell and insoluble substrate or though mediated processes
involving shuttles that move electrons to and from the microbial cell to an insoluble substrate.
Cyclic voltammetry tests were conducted on biofilms, planktonic cells, and filtered spent media
to assay for electrochemical activity in each fraction (Figure 3.11, Appendix S1). Analysis of CV
plots revealed what fractions contributed to electrochemical activity by identifying peaks
suggestive of oxidation or reduction (Figure 3.11). Midpoint potentials of the redox active peaks
were calculated (Table 3.4). A diverse range of midpoint potentials was obtained, with the iron
oxidizing isolates possessing more reducing midpoint potentials (-66.86 to -389.1 mV vs.
Ag/AgCl) compared to sulfur oxidizing isolates (99.20 to -59.4 mV vs. Ag/AgCl).
Electrochemical activity was determined to come from biofilm/attached cells for all isolates.
SEM imaging of electrodes revealed attached cells and biofilms on the electrode materials
(Figure 3.13).

Flavin production by the isolates was determined by running filtered samples taken from
electrochemical cells during chronoamperometry tests through HPLC. Flavins were detected in
79
the nM range for all the Gammaproteobacteria strains isolated (Figure 3.12). No flavins were
detected in the samples from other strains. Both Marinobacter strains produced FMN at a similar
level (~40-50 nM). Although every isolate consistently produced FMN at some level, there was
variability in the detection of RF. Samples taken from the electrochemical cells were taken at
different time points during chronoamperometry experiments. These experiments should be
repeated with consistent sampling time points at specific growth phases to further understand the
absence and presence of RF. These data suggest endogeneous shuttles are produced consistently
in the form of FMN for all Gammaproteobacteria strains. Interestingly, CV tests on filtered
spent media did not indicate the presence of redox active shuttles. Flavins are known to bind to

Figure 3.12. Flavin production of the Gammaproteobacteria isolates. Flavin mononucleotide
concentrations (FMN) are indicated by the red bars and riboflavin (RF) concentrations are
represented by the blue bars. Bars are the mean and s.d. of at least triplicate samples for FMN.
Bars are the mean and s.d. of at least duplicate samples for RF, with the exception of the RF
value for -500 mV Isolate 1. RF was only detected once for -500 mV Isolate 1.

80
filters, which might hinder their detection. Since all isolates exhibited electrochemical activity
when attached to electrodes, the role of the flavins detected might be to bind to outermembrane
proteins to facilitate EET rather than act as shuttles themselves.

Figure 3.13. SEM images of the isolates. A) -550 mV Isolate 1 (Vibrio sp.) grown on a
carbon felt electrode and B) -650 mV Isolate 1 (Marinobacter sp.) grown on a carbon felt
electrode.

DISCUSSION

Our secondary bioreactor electrochemical enrichments allowed simultaneous simplification of
the system to enrich for cathode-oxidizing microorganisms and the ability to continuously
monitor electron uptake activity. The addition of a selected TEA (sulfate or nitrate) to the
secondary bioreactors was coupled to negative current generation indicative of uptake of
electrons by the cells from the cathode (Figure 3.2). For the sulfate fed secondary bioreactors
(-550, -650, and -750 mV), the enriched electrode biofilm communities showed a higher
abundance of Deltaproteobacteria compared to the planktonic communities (Figure 3.5A).
Deltaproteobacteria have been shown to be strongly associated with anode electrodes compared
to solutions in MFC systems (Ishii et al. 2014). The predominant mechanisms of EET
characterized in Deltaproteobacteria involve direct contact between the microbial cell and
B) -650 mV Isolate 1 A) -550 mV Isolate 1
81
insoluble substrate (Lovley et al. 2004; Logan 2009). The predominance of Deltaproteobacteria
in electrode communities was not observed for the -400 mV secondary bioreactors, which
utilized nitrate as the primary TEA. This is not surprising as the choice of TEA influenced the
resultant community structures. Electrode biofilm communities from the nitrate fed secondary
bioreactor (-400 mV) exhibited significant enrichments of the Rhodobacterales and
Chromatiales not seen in the sulfate fed secondary bioreactors. The applied redox potential
(electron donor source) was the other primary driving factor of bioreactor community structure.
The -550 mV bioreactor communities showed greater similarity to the -400 and -500 mV
bioreactor communities although they differed in TEA (Figure 3.6). This similarity was further
corroborated by the increased abundance of nitrate reductase in the -550 mV bioreactor
communities compared to the other sulfate fed secondary bioreactors (-650 and -750 mV). This
could be explained by the fact that the -550 mV redox potential was not as favorable
energetically to be coupled to sulfate reduction, and thus the enriched microbes were more
versatile in what electron acceptors could be used.

The diverse starting redox potentials utilized resulted in the subsequent isolation of six new
electrochemically active strains. It is important to note that all of the isolates obtained fall within
the major groups that were enriched during secondary electrochemical bioreactor operation
(Figure 3.5, Table 3.4). Our laboratory’s initial study resulted in the successful isolation of
several electrochemically active strains in the Alphaproteobacteria and Gammaproteobacteria
(Rowe et al. 2015). With this work, we have expanded the diversity of isolates into the
Epsilonproteobacteria, Clostridia, and Deltaproteobacteria. An Arcobacter strain, -750 mV
Isolate 1 was obtained through our sulfur-oxidizing tertiary enrichments. Recently, several
Arcobacter OTUs were found to colonize iron sulfide minerals incubated around Catalina Island
82
(Ramírez et al. 2016). To this date, electrochemical activity has only been described in one
Arcobacter strain, Arcobacter butzleri ED-1, which is capable of anode reduction (Fedorovich et
al. 2009). This strain was isolated from a microbial fuel cell inoculated with marine sediment,
and a proteomic investigation revealed differential expression in FlaA, a flagellin and two novel
putative c-type cytochromes in anode associated cells compared to cells grown in normal batch
culture (Pereira-Medrano et al. 2013). There is much to be elucidated in how members of the
Arcobacter genus perform EET as we are just beginning to obtain electrochemically active
isolates within the Epsilonproteobacteria.

In this study, we report the first instance of an electrochemically active isolate from the genus
Vallitalea (-750 mV Isolate 2, Vallitalea sp.). There have been several reports of other members
of the Clostridia exhibiting EET activity (Kopke et al. 2010; Wrighton et al. 2011; Choi et al.
2012; Choi et al. 2014), but the Vallitalea are a poorly described genus with no reports of
electrochemical activity. The two recognized species of the genus were both isolated from
hydrothermal systems, marine sediments from the Guaymas basin (Lakhal et al. 2013) and a
chimney from Prony Bay in New Caledonia (Aissa et al. 2014). Further characterization of the
Vallitalea isolate is essential to expand understanding of this genus and its potential for EET as it
is commonly found in various environments (Catania et al. 2016, Pootakham et al. 2017). One
isolate from the Deltaproteobacteria was obtained, -550 mV Isolate 2 (Desulfovibrio sp.).
Members of the Desulfovibrio genus are found in a wide variety of environments and are
characterized as sulfate reducers (Garrity et al. 2005). Although the direct role of sulfate
reducing bacteria (SRBs) in metal corrosion is a controversial topic, there have been studies that
suggest SRBs are capable of directly taking up electrons from iron (Dinh et al. 2004; Enning et
al. 2012; Venzlaff et al. 2013). Some of the precipitates formed on the elemental iron used in
83
tertiary enrichment cultures (Figure 3.8C) resembled the mineral crust (FeS, FeCO
3
, Mg/CaCO
3
)
described as being formed as a result of electron uptake from iron by these SRBs (Enning et al.
2012; Venzlaff et al. 2013). In addition, in a study screening for cathode oxidation ability in
several environmental isolates, production of cathodic current by Desulfovibrio piger was
demonstrated (deCamposRodrigues & Rosenbaum 2014). The mechanisms of EET in SRBs are
currently unknown, although potential proteins involved have been suggested through
metagenomic studies (Skennerton et al. 2017).

The two Marinobacter strains isolated came from different sources (-500 mV and -650 mV
secondary bioreactors) and different subsequent tertiary electron donors (sulfur and iron,
respectively). They also exhibited very different midpoint potentials, with the iron-oxidizing
isolate possessing a significantly more reducing midpoint potential (-389.1 mV) compared to the
sulfur-oxidizing isolate (35.09 mV). The two Marinobacter strains from our initial study were
both isolated from FeS and had midpoint potentials between -176.2 and -184 mV (Rowe et al.
2015). These results suggest great metabolic flexibility and potentially diverse mechanisms of
EET in the Marinobacter genus. The Marinobacter genus is ubiquitous in marine environments,
and some Marinobacter spp. are known neutrophilic Fe(II) oxidizers (Edwards et al. 2003; Bonis
& Gralnick 2015). Genomic analysis of Marinobacter aquaeolei VT8 revealed the presence of
47 genes encoding cytochrome proteins (Singer et al. 2011). Many bacteria utilize
outermembrane cytochromes to carry out EET (Myers & Myers 1992; Kim et al. 2006).
Additionally, a member of the Marinobacter was shown to be a predominant constituent of a
biocathode derived from marine sediment (Strycharz-Glaven et al. 2013). The Marinobacter
strains our laboratory has isolated are prime candidates to further investigate the mechanisms of
cathode oxidation in this metabolically versatile genus. A Vibrio strain, -550 mV Isolate 1, was
84
another Gammaproteobacteria sulfur-oxidizing isolate obtained. Electrochemical activity in the
Vibrio genus has not been widely reported, although there was a study that demonstrated Vibrio
parahaemolyticus, a pathogen with an MtrB (outermembrane protein identified in Shewanella
oneidensis MR-1) homolog was capable of reducing insoluble iron and manganese (Wee et al.
2014). With our Vibrio isolate, anodic current was demonstrated before switching over to
cathodic conditions suggesting the strain is also capable of electrode reduction (Figure 3.10). The
observed homology between the two strains of Gammaproteobacteria to perform metal
reduction, provides a starting point to investigate EET in our Gammaproteobacteria isolates.

The electrochemical characterization of the isolates highlighted the diversity in the dominant
redox potential of the proteins that interact with the electrode (demonstrated by midpoint
potentials) (Table 3.4). Generally, a trend towards more reducing midpoint potentials for iron-
oxidizing isolates (-66.86 to -389.1 mV) compared to sulfur-oxidizing isolates (-59.40 to 99.20
mV) was observed and is consistent with results from our initial study (Rowe et al. 2015). All of
our isolates demonstrated electrochemical activity in the attached/biofilm phase. The detection of
flavins in both the secondary bioreactors (Table 3.2) and from isolate chronoamperometry tests
(Figure 3.12) suggests that they play a role in cathode oxidation. Further investigation is needed
in the potential involvement of flavins in electrode oxidation for our Gammaproteobacteria
isolates. Although our isolations captured more diversity in the type of genera obtained, we still
did not attain isolates from groups that seemed to contribute to cathode oxidation in the
secondary bioreactors. For example, the Chromatiales were highly enriched in the -400 and -500
mV secondary bioreactor communities, and were shown to contribute the most to the presence of
nitrate reductase (Figure 3.7). Further work will look at using electrochemical isolation
techniques to target groups of interest rather than traditional isolation techniques.
85
CONCLUSION

Our work using five different redox potentials was able to enrich for six novel electrochemically
active isolates from one environmental system. These isolates have further expanded the
diversity of microorganisms capable of cathode oxidation. It is becoming more evident that
microorganisms that are able to perform anode reduction are also capable of cathode oxidation
(Gregory et al. 2004; Rosenbaum et al. 2011; Rowe et al. 2017). Two of the isolates obtained in
this study come from genera (Arcobacter and Vibrio) where insoluble substrate reduction has
been shown with no record of cathode oxidation. Screening bacteria that are traditionally known
as metal reducers for cathode oxidation might reveal that the flexibility to also oxidize insoluble
substrates is relatively common and explain the prevalence of some of these organisms in
oligotrophic environments. Expanding electrochemical techniques to probe environments where
reduced minerals predominate will help further the understanding of these metabolisms.
Insoluble substrate oxidation is an ability that seems to be widespread, and we are only
beginning to understand the extent of this process in nature.

86
APPENDICES

Appendix S1. Representative cyclic voltammetry plots under non-turnover
conditions for isolates listed (A-D). Plots of CVs conducted on biofilm, planktonic, and
control sterile artificial seawater (ASW control). Scans were run at 5 mV/sec.

87
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CHAPTER 4

DETECTING MICROBIAL METABOLISM AND IDENTIFYING BIOSIGNATURES IN
SOILS WITH BIOELECTROCHEMICAL DEVICES

INTRODUCTION

The first attempt to detect microbial life and metabolism on Mars was through the biological
experiments conducted on the Viking mission in 1976. These experiments focused on analysis of
organic chemical signatures and metabolic activity determined through the consumption and
transformation of introduced organic compounds (Klein 1978; Brown et al. 1978; Levin and
Straat 1979). Results from these types of analyses must be interpreted with caution due to
difficulties of distinguishing biotic chemical signatures from similar signatures produced by
abiotic processes. The positive results from the Viking biological experiments were attributed to
the oxidizing agents produced by the ionizing radiation of perchlorate (Quinn et al. 2013).
Additionally, gas chromatography-mass spectrometry analysis was unable to detect any organic
compounds in the soil samples (Klein 1978). After the overall negative conclusion of the
metabolism-based investigations of the Viking mission, life-detection instrument development
has been biased towards looking for what is known on Earth in the form of biomolecules such as
amino acid abundances and specific organic structures like lipids as components of biomass
(Nealson & Conrad 1999; Conrad and Nealson 2001; Dorn et al. 2003). Habitability is often
defined more broadly in a “follow the energy” approach that searches for useful redox couples in
the environment (NAI 2015; Chyba & Phillips 2001; Johnson et al. 2003; Hand et al. 2007;
Kounaves et al. 2009; Grotzinger et al. 2012; Blake et al. 2012) to identify likely inorganic and
organic components that could provide energy for metabolism. The identification of habitable or
formerly habitable environments on other worlds (Greenberg et al. 2000; Parkinson et al. 2007;
95
Parkinson et al. 2008; Stoker et al. 2010; Grotzinger et al. 2014) has provided us with sites to
start looking for life with a strategy that builds upon what we have learned from former missions
(e.g. the Viking mission) but is also novel in its approach. To overcome this challenge, and be
capable of detecting life with a different origin than on Earth, focus must be placed on designing
instruments that are capable of detecting a biosignature that is a shared and distinguishable
feature of all living organisms while being as non-Earth centric as possible (Conrad & Nealson
2001). To this end, we are working towards developing and implementing BioElectroChemical
Devices (BECDs) as life-detection instruments to measure microbial metabolism in habitable
environments.

Metabolism is a universal characteristic of (earthly) life involving electron flow (redox) reactions
among organic and/or inorganic substrates. All metabolic modes use the same basic processes of
differential redox and electron transfer to convert energy from the environment into biologically
functional energy in the form of adenosine triphosphate (ATP) (Guzman & Martin 2009;
Srinivasan & Morowitz 2009). This process involves oxidation of an electron donor (fuel),
liberating electrons and protons. The electrons are eventually transferred to an electron acceptor
(oxidant) reducing it in the process. The electron flow from an electron donor to an electron
acceptor results in protons being transported across a cellular membrane to produce an
electrochemical gradient that drives the synthesis of ATP, the energy currency used for all
cellular processes.

Since metabolism always involves redox reactions, it can be detected through either chemical or
electrical means. Changes in oxidation state in a reaction can be analyzed using electrochemical
techniques such as fuel cells, which measure electrical current produced by oxidation and
96
reduction half-reactions occurring in separate reservoirs divided by a proton exchange
membrane. In a fuel cell, the electrodes (where oxidation or reduction occurs) are often coated in
a catalyst (e.g. platinum or other metals) to increase surface area and lower the energy barrier for
the oxidation or reduction reaction. BECDs operate similarly to standard fuel cells, but instead of
an inorganic metal catalyst in the anode compartment, microorganisms catalyze the oxidation
reactions. In the anode compartment, microorganisms oxidize an electron donor, releasing both
electrons and protons. These electrons are directly transferred to the anode, and subsequently
travel through an external conductor (e.g. a resistor) to reach the cathode compartment.
Reduction reactions (often the reduction of O
2
to H
2
O) occur at the cathode, utilizing the
electrons and the protons that were released. When an external conductor is placed between the
anode and cathode, current is produced and an increase in electrical potential can be measured.
The current produced is proportional to the oxidative processes occurring in the anode
compartment, either from microbial metabolism or other redox reactions. The physiological
process of transferring electrons directly to an electrode is known as extracellular electron
transport (EET). Though once thought to be rare, EET capability has been found in a wide
variety of environmental microbial communities (Holmes et al. 2004; Wrighton et al. 2010; Ishii
et al. 2012) including those in extreme environments (Miller & Oremland 2008; Rojas et al.
2017) and is a biological process that should be considered when looking for extraterrestrial life.

The diversity of microbial metabolism is vast, as microbes are able to use a variety of substrates
as electron donors and acceptors as long as it is thermodynamically favorable for energy to be
obtained. Both electron donors and acceptors can take various forms, e.g. a diffusible soluble
substrate (lactate as an electron donor), diffusible gas (H
2
as an electron donor), or even a solid
conductive mineral or electrode (electron donor or acceptor) for microbes that are capable of
97
EET. Different species and strains of microbes perform specific redox reactions that utilize
particular electron donors or acceptors, and these occur at specific potentials. BECDs can also be
configured as a three-electrode system, where the working electrode can be poised at a biological
redox potential of interest. The redox potential can be controlled so that the electrode can serve
as either an electron donor or acceptor, and current at the working electrode can be measured and
is indicative of active redox reactions. Consequently, the electrical current generated by
microbial metabolism can be identified in-situ by placing electrodes directly in the environment
or screened in samples ex-situ. Environmental samples from inhabited niches should contain all
the components necessary to produce current flow, i.e., catalysts (microorganisms) and fuel
(nutrients). BECDs can also probe for starving and/or dormant microorganisms by artificially
adding a fuel to drive metabolism and/or growth. A previous study suggested microbial fuel cells
as a strategy for life detection by successfully measuring the potential generated by monocultures
of microorganisms and by the microbial community present in a humus-rich soil (Abrevaya et al.
2010).

In this paper, we demonstrate that BECDs can detect microbial metabolism, and that the
sensitivity of BECDs can be enhanced by adding fuel (organic substrates) to an existing
environmental sample. BECDs improve upon the detection of metabolism approach utilized by
the Viking missions by looking for temporal patterns indicative of metabolism as well as
identifying electrical signatures that demonstrate active metabolism rather than the chemical
(waste) byproducts of metabolism. The use of two different soil mixtures, a commercially
available soil and a Mars simulant soil, offers a broader perspective on what electrochemical
signals can be produced abiotically and perhaps biotically. Additionally, by using a model
98
organism, Shewanella oneidensis MR-1, we were able to understand how varying the starting
number of cells affects the electrochemical signal produced.

MATERIALS & METHODS

BECD experiments were performed using microbial fuel cell (MFC) devices that contain two
separated reservoirs across a proton exchange membrane (Figure 4.1A) or a three-electrode
electrochemical cell with a working electrode, reference electrode, and counter electrode (Figure
4.1B). For MFC experiments, one reservoir contained the anode immersed in a volume of wet
soil (soil and anolyte) and the other reservoir contained the cathode immersed in the catholyte
solution (Table 4.1). MFC experiments either used unsterilized soil or sterilized soil for controls.
Injections of a fuel (electron donor in the form of lactate) for microbial growth were added at
periodic intervals. Voltage was measured and current was calculated between the two electrodes
to determine the presence of microbial metabolism. In three-electrode electrochemical cell
experiments, a single compartment contained a working electrode, reference electrode, and
counter electrode immersed in defined minimal media with lactate as an electron donor. The
working electrode was poised at a redox potential that allowed for it to be used as an electron
acceptor by S. oneidensis MR-1. Different inoculums in terms of cell number were injected into
the three-electrode electrochemical cells. Potential at the working electrode was controlled by a
potentiostat, and the current produced at the working electrode was continually measured to
determine the rate at which S. oneidensis MR-1 cells were actively transferring electrons to the
working electrode.

Table 4.1. Summary of substrates used in BECD experiments.
99

Soil, Anolyte and Catholyte Preparation for MFC Experiments

The soil mix in the anode compartment contained either standard potting soil, or Mars simulant
soil. Standard potting soil mix (referred to here as HF, Happy Frog brand) was procured from
FoxFarm Soil & Fertilizer Company, Samoa, CA. The HF soil was chosen compared to other
commercial potting soils, because it omits standard pre-sterilization procedures that eliminate the
endemic microbial communities present in soils. Mars simulant soil (referred to here as MSS)
was JSC-Mars-1A (M-JSC1A; Orbitec, Madison, WI), which is accepted as a spectral and
physical simulant for testing various instruments and hardware in preparation for Mars missions
(Hudson et al. 2007; Chevrier et al. 2007; Pommerol et al. 2009). JSC-Mars-1A soil simulant has
been characterized in detail (Allen et al. 1998), and it is sourced from the Pu’u Nene cinder cone
in Hawaii. Soil samples were divided into 20 g batches and placed in sterile, acid-washed flasks.
Some were sterilized by autoclaving with two cycles: one cycle at 121°C for 1 hr and 45 min and
a second cycle with the same conditions 24 hr after the first cycle. Other samples of MSS and HF
potting soil were not sterilized and were used as is. Soils (sterile and non-sterile) were hydrated
up to 30 mL total volume with anolyte solutions. Sterile 0.6 M NaCl was used as the anolyte for
all HF potting soil experiments. Sterile Mars simulant salt solution was used as the anolyte for all
Mars simulant soil experiments. The Mars simulant salt solution contained: 1.2 mM Mg(ClO
4
)
2
,
1.8 mM MgSO
4
, 3.6 mM CaSO
4
, 0.3 mM KCl, 89.7 µM NaCl, 0.6 mM Na
2
CO
3
, and 20 mM
CaCO
3
, and is derived from a mixture of salts detected at the Mars Phoenix mission landing site
100
(Hecht et al. 2009). The sterile catholyte solution contained 50 mM potassium ferricyanide
(K
3
Fe(CN)
6
) and 50 mM K
2
HPO
4
(pH = 7.2). Potassium ferricyanide was chosen as a catholyte
as it does not require O
2
for cathodic reactions to proceed and would function in oxygen free
atmospheres as part of a Mars instrument. The fuel (electron donor) injection solution contained
50 mM lactate, 50 mM piperazine-N,N’-bis(2-ethanesulfonic acid) (PIPES) buffer and was
prepared using traditional anaerobic techniques (Miller & Wolin 1974). 50 mM PIPES buffer
solution was also prepared as a control for fuel addition experiments to account for responses to
fuel injection being due to the PIPES buffer and not the lactate (electron donor).

Minimal Media, Cell Culturing, and Inoculum Preparation for Three-Electrode Chemical
Cell Experiments
S. oneidensis MR-1 is a gram-negative, facultative anaerobic bacterium capable of using a wide
variety of electron acceptors including mineral oxides through EET (Myers & Nealson 1988;
Nealson & Saffarini 1994; Heidelberg et al. 2002). M1 minimal media for S. oneidensis MR-1
experiments contained: 50 mM PIPES buffer, 7.5 mM NaOH, 28.04 mM NH
4
Cl, 1.34 mM KCl,
4.35 mM NaH
2
PO
4
⋅H
2
O, and 18 mM C
3
H
5
O
3
Na (lactate). Lactate served as the electron donor
for all experiments. 10X minerals, vitamins, and amino acid solutions were also added to the
minimal media as previously described (Bretschger et al. 2007). Luria Broth (LB) agar plates
were streaked with frozen stock of S. oneidensis MR-1. A single colony was selected from LB
agar plates and inoculated into 50 mL of liquid LB. After 24 hours, 50 µl of the liquid culture
was transferred to 50 mL of M1 minimal media. After 24 hours, the optical density (OD) of the
culture was taken at 600 nm and adjusted to a desired cell density using 50 mM PIPES buffer.
All culturing steps were conducted at 30°C.

101

Figure 4.1. General schematic of the BECD systems used, A) MFC system and B) three-
electrode electrochemical cell. For MFC systems, the anode and cathode chambers were
separated by a proton exchange membrane. Anode compartments contained the soil hydrated in
anolyte solution while the cathode compartments contained the catholyte (50 mM K
3
Fe(CN)
6
, 50
mM K
2
HPO
4
). For three-electrode systems, the working electrode (anode) potential was
controlled by a potentiostat. Three-electrode electrochemical cells contained minimal media.

A)
B)
102
BECD Experiments

MFC Experiments: MFC cells consisted of dual compartment 30 mL custom designed glass
chambers (Figure 4.1). Anode and cathode chambers were separated by a pretreated Nafion 424
ion exchange membrane (DuPont, Wilmington, DE). Briefly, ion exchange membranes were
soaked in 0.5% HCl and rinsed with ultrapure H
2
O, and subsequently boiled in 3% H
2
O
2
and 0.5
M H
2
SO
4
with ultrapure H
2
O wash steps in between and after each boiling treatment. All
electrodes were made from 6 mm thick graphite felt (Electrolytica, Alpharetta, GA) cut into
circles with a 39 mm diameter and treated as previously described (Hsu et al. 2012). Graphite felt
was bonded to platinum wire (Alfa Aesar, Tewskbury, MA) with EPOX-4 graphite adhesive
(Electrolytica, Alpharetta, GA). The anode chamber was continuously purged with filtered N
2
and the cathode chamber was continuously purged with filtered air at a rate of 15 mL/min.
Voltage across a 10 Ω resistor was measured and recorded every 5 minutes by a high-impedance
digital multimeter (Keithley, Cleveland, OH). For organic acid (electron donor) stimulation
experiments, fuel cells were run for 24 hr to collect baseline voltages. Lactate (3 mM final
concentration) or PIPES buffer were injected into the anode compartments after 24 hr and two
additional injections were performed at the 48 hr and 72 hr mark. Current (I) was calculated
according to I=V/R, and current density (j) was determined by dividing I by the surface area of
the electrode (20 cm
2
).

Three-Electrode Electrochemical Cell Experiments: Tests were performed in the same chambers
as in the fuel cell experiments configured to have a port for a 1M KCl Ag/AgCl reference
electrode (BASi, West Lafayette, IN). Graphite felt electrodes (as described above) served as the
working electrode, and platinum wire was used as a counter electrode. The three-electrode
103
electrochemical cells were continuously purged with filtered N
2
. The working electrodes were
poised at an electron accepting redox potential of +400 mV (vs. Ag/AgCl) using a Quadstat
potentiostat (eDAQ, Colorado Springs, CO). Current was measured and recorded at the working
electrode every minute using the eCorder eCHART software every minute (eDAQ Inc., Colorado
Springs, CO).

Cyclic Voltammetry of Test Substrates

Cyclic voltammetry (CV) tests were conducted on both hydrated soils (sterile and non-sterile)
and anolyte solutions to observe electron transfer reactions (oxidation and reduction peaks) of
the samples. Tests were performed with the same set-up and three electrodes as described above
for the three-electrode electrochemical cell. The three-electrode chamber was purged with
filtered N
2
. CVs were performed with a Gamry Reference 600 potentiostat and Gamry
Framework software (Gamry Instruments, Warminster, PA). Each CV analysis was conducted at
a scan rate of 10 mV/sec over a range of -1000 to 1000 mV vs. Ag/AgCl for 3 cycles.

Analysis of Organic Acids and Proteins

Samples taken from the anode chambers were filtered through 0.2 µm Acrodisc Supor®
membrane syringe filters (Pall Corporation, Port Washington, NY) for organic acid analysis.
Samples were stored at -80°C until analysis. 2 mL aliquots of removed soil anolyte were
centrifuged at 13,000 rpm for 5 min to concentrate cells for total protein analysis. Pellets were
stored at -20°C until analysis. After fuel cell experiments were done, electrodes were sectioned
in half to conduct both total proteins quantification and scanning electron microscopy.

Organic acid concentrations were measured as described previously (Bretschger et al. 2010) by
using a high-performance liquid chromatograph (HPLC) with a Hi-Plex PLRP-S 300 mm x 7
104
mm column (Agilent Technologies, Santa Clara, CA). A fluorescence detector (Agilent
Technologies, Santa Clara CA) was used with an excitation wavelength of 450 nm and an
emission wavelength of 520 nm. The HPLC was operated with 0.1 M sulfuric acid as the mobile
phase (0.5 mL/min). To measure total proteins, pellets were hydrolyzed in 1 M NaOH at 50°C
for 30 min with vigorous mixing by vortex at 10 min intervals. Electrode samples were
hydrolyzed in 200 mM NaOH at 100°C for 90 min with vigorous mixing by vortex at 15 min
intervals. Soluble protein in the extracts was determined spectrophotometrically using Folin
phenol reagent (Lowry et al. 1951) with bovine serum albumin (BSA) as a standard.

Scanning Electron Microscopy (SEM) Sample Preparation of Electrode Samples

Electrode samples were fixed in distillation purified electron microscopy grade 2.5%
glutaraldehyde (Electron Microscopy Sciences, Hatfield, PA). Samples were then processed
through an ethanol dehydration series (30, 50, 70, 80, 90, 95, 100% v:v ethanol, three 10 min
intervals for each concentration) and critical point drying (Autosamdri 815 critical point drier,
Tousimis Inc., Rockville, MD). Samples were mounted on aluminum stubs and coated with Au
(Sputter Coater 108, Cressington Scientific, Watford UK). Images were captured at 5 keV using
a JOEL JSM 7100F low vacuum field emission scanning electron microscope (JEOL USA, Inc.,
Pleasanton, CA)

RESULTS
Cyclic Voltammetry of Test Substrates: Cyclic voltammetry (CV) was conducted by scanning
the different soil substrates and anolytes across a voltage range of -1000 to 1000 mV vs.
Ag/AgCl. CV analysis was used to detect different oxidation and reduction reactions in the test
substrates. For both soils, commercial potting soil (HF) and Mars simulant soil (MSS), the
105
sterilized soils showed much more activity in terms of reduction and oxidation reactions (Figure
4.2). For the HF soil, there were no responses observed in the non-sterile soil, while the sterile
soil exhibited substantial anodic (reductive) current at +250 mV and cathodic (oxidative) current
at -250 mV. The MSS soil possessed a similar trend, with the sterile soil showing pronounced
anodic current at +500 mV and cathodic current at -250 mV. This data suggests that the
sterilization process of autoclaving releases redox active species in both soils.

Figure 4.2. Cyclic voltammograms of the various experimental test substrates: A) Sterile
Happy Frog Soil, Non-Sterile Happy Frog Soil, and 0.6 M NaCl, and B) Sterile Mars Simulant
Soil, Non-Sterile Mars Simulant Soil, and Mars Simulant Salt Solution.

Long-term baseline fuel cell tests: Endemic microbial metabolic activity of both soils was
investigated with long-term fuel cell experiments. Sterile and non-sterile soils were hydrated and
placed in a standard MFC set-up (Figure 4.1B). Long-term fuel cell tests were run for 336 hr
after preliminary tests (Appendix S1) indicated a longer run time was necessary to allow for
current density readings to stabilize. Non-sterile HF soil exhibited an increased amount of
current density than sterile HF soil (Figure 4.3A). The delineation between the sterile and non-
sterile HF soil occurred after 80 hr. With the long-term fuel cells tests for the MSS soil, the
106
sterile MSS soil had a higher current density than the non-sterile MSS soil starting at 25 hr
(Figure 4.3B). There was a large increase in current density seen in the sterile MSS soil that
began at the 25 hr mark, which eventually started to decay after 30 hours.

Figure 4.3. Long-term (2-week) baseline full cell tests conducted on: A) Happy Frog Soil, B)
Mars Simulant Soil.

Fuel cell tests with organic acid addition: Organic acid addition to both soils was explored as a
way to enhance microbial metabolic activity and observe the subsequent responses in terms of
current density. Lactate was chosen to serve as the organic acid as it is a common monovalent
organic acid found in soils that is weakly adsorbed to the solid phase (Jones et al. 2003). In
addition, lactate is a byproduct of complex carbon substrate oxidation that can be utilized as an
electron donor by many microbial groups, and has been shown to increase metal reduction rates
of anaerobic microbial communities in situ (Brodie et al. 2006; Faybishenko et al. 2008;
Burkhardt et al. 2010). Lactate injections in replicate fuel cell experiments with HF soil,
displayed a sustained and increasing current density in the non-sterile soil (Figure 4.4A). The
control experiments, non-sterile soil with PIPES addition and sterile soil with lactate addition,
107
demonstrated a spike in current density with injections that decays and is not sustained with time.
Similar trends were also observed in the lactate addition experiments done on MSS Soil, with the
exception of the sterile MSS with lactate addition (Figure 4.5). The non-sterile MSS soil had an

Figure 4.4. Fuel cell experiments with organic acid additions conducted on HF Soil. Each
arrow corresponds to either a 3 mM lactate or PIPES addition. A) Replicate non-sterile HF with
lactate addition experiments, B) Replicate sterile HF with lactate addition experiments, C)
Replicate non-sterile HF with PIPES addition experiments, and D) Average of all three
conditions, dashed lined indicates maximum current density achieved in non-sterile long-term
fuel experiments with no organic addition.

increasing amount of current density with each lactate injection (Figure 4.5A). The sterile MSS
soil replicate experiments with lactate injections, also showed a gradual increase in current, but
the readings never reached the level of the non-sterile MSS soil. The non-sterile MSS soil
experiments exhibited an increase in current density after the first PIPES injection, but the
current dropped off during the time course of the rest of the experiments. The maximum stable
108
level of average current obtained for the non-sterile MSS soil was 3.77 nA/cm
2
and 1.97 nA/cm
2

for the non-sterile HF soil.

Figure 4.5. Fuel cell experiments with organic acid additions conducted on MS Soil. Each
arrow corresponds to either a 3 mM lactate or PIPES addition. A) Replicate non-sterile MSS
with lactate addition experiments, B) Replicate sterile MSS with lactate addition experiments, C)
Replicate non-sterile MSS with PIPES addition experiments, and D) Average of all three
conditions, dashed line indicates maximum current density achieved in non-sterile long-term fuel
cell experiments with no organic addition.

Analysis of organic acids and protein concentrations during fuel cell experiments with
organic acid addition: Samples were taken during the time course of the organic acid addition
fuel cell experiments to measure organic acid (lactate, acetate, and pyruvate) concentrations.
Acetate and pyruvate were measured as they are both byproducts of lactate oxidation. At each
injection time point, 1.5 mL of a 50 mM lactate solution was injected into the anode
compartment. Table 4.2 summarizes the measured organic acid concentrations for a
representative set of the lactate addition experiments done on each soil. Accounting for all of the
109
lactate additions, a final concentration of ~9 mM lactate would be expected if all of it remained
in solution. For both soils, the sterile experiments showed an incremental increase in lactate
concentrations. At the end of the experiments, 3.11 mM lactate was detected in the non-sterile
HF soil and 4.61 mM lactate was detected in the non-sterile MSS soil. An incremental increase
in pyruvate was observed in the non-sterile HF soil experiment. For the non-sterile MSS soil
experiment, acetate was only observed at the end of the experiment.

Table 4.2. Concentrations of lactate and potential byproducts from lactate oxidation,
acetate and pyruvate, during the time course of representative fuel cell experiments with
lactate addition. Time points, T1, T2, and T3 occurred before ~3 mM lactate injections. T4 was
the final time point at experiment end.

ND indicates not detected

Total proteins were measured in samples taken from the anode compartment to evaluate changes
in biomass during the time course of the organic acid addition experiments (Figure 4.6). For the
HF soil experiments, the initial protein concentrations across all experimental conditions were in
a similar range. The sterile HF soil experiments displayed a decrease in protein concentrations
with each subsequent sample, with the exception of the final time point that displayed a slightly
elevated protein concentration. The non-sterile HF soil experiments with PIPES injections
exhibited decreases in protein concentration during the experimental time course. In the non-
sterile with lactate addition replicate experiments, protein concentrations remained constant or
110
increased slightly during the initial time points until dropping off at the T3 time point. For the
MSS experiments, protein concentrations in the non-sterile lactate addition experiments had
markedly higher protein concentrations initially and throughout the time course of the
experiments compared to the other conditions. In the non-sterile MSS soil PIPES addition
replicate experiments, protein concentrations generally remained constant. With the sterile MSS
soil lactate addition experiments a decrease in protein concentrations is observed during the time
course after an initial increase at the T1 time point. Protein concentrations from the electrodes
were also measured to account for any biomass that was attached to the electrode. For both soils,

Figure 4.6. Average protein concentrations from soil anolyte samples taken during the time
course (T0: experiment start to T4: experiment end) of replicate fuel cell with organic acid
(lactate) addition experiments. The protein concentrations measured reflect any biomass in the
planktonic phase in the soil anolyte.

111
the experimental conditions with lactate had similar values in protein concentrations, while the
PIPES addition experiments had markedly less protein concentrations (Figure 4.7). Protein
samples taken from the anode compartment were generally higher in the MSS soil experiments
compared to HF soil experiments (Figure 4.6), while the reverse is true for the electrode protein
concentrations (Figure 4.7). Cell biomass attached to the electrodes was also evaluated with
SEM imaging (Appendix S2). No dense biofilms were detected, although cells were observed
attached to the electrodes from non-sterile soil experiments.

Figure 4.7. Average protein concentration from electrodes harvested after replicate fuel
cell with organic acid (lactate) addition experiments. The protein concentrations measured
reflect any biomass that attached to the electrode.

Three-electrode system experiments with different cell inoculums: Experiments using a three-
electrode BECD system were designed to evaluate the minimal amount of biomass (in terms of
cell number) needed to detect an electrochemical signal. The current density generated with
112
different initial cell numbers of S. oneidensis MR-1 is shown in Figure 4.8. All inoculums used
(1.06 x 10
5
to 7.68 x 10
6
cells/ml) exhibited an increase in current density after 25 hr. The
maximum amount of current density (2.98 to 4.78 µA/cm
2
) was similar for all of the cell
inoculums in the 10
6
cells/ml range. The maximum amount of current density achieved was
22.15 µA/cm
2
with the 2.07 x 10
5
cells/ml inoculum. The 1.06 x 10
5
cells/ml inoculum also
generated a higher current density (11.89 µA/cm
2
) when compared to the experiments where
higher initial cell numbers were used.

Figure 4.8. Current density generated in three-electrode electrochemical cells inoculated
with differing cell inoculums of S. oneidensis MR-1.

113
DISCUSSION
BECD systems such as MFCs have been suggested to be a viable method for the in situ detection
of microbial life (Abrevaya et al. 2010). This study demonstrated the viability of MFCs to detect
microbial metabolism with pure cultures of Saccharomyces cerevisae and Natrialba magadii and
in one environmental sample of humus-rich topsoil (Abrevaya et al. 2010). The detection of life
will likely not occur in environments that are optimized for the growth of one organism or in
environments that are replete in organics. Microorganisms found in the most oligotrophic
subsurface environments on Earth are estimated to catabolize 10
4
to 10
6
more slowly than
organisms grown in the laboratory, with biomass turnover occurring at timescales of hundreds to
thousands of years (Hoehler & Jørgensen 2013b). In our study, we chose to explore the efficacy
of BECDs for life detection by using two different soils that were not altered by the addition of
nutrients, including a Mars simulant soil. In long-term MFC tests, there was minimal
differentiation between sterile and non-sterile HF soil, and high variability in the response of
MSS soil (Figure 4.3, Appendix S1). These results suggest simple hydration of the soils was not
enough to stimulate metabolic activity, and differentiate between electrochemical signals
produced biotically or abiotically.

Many microorganisms in the natural environment are not actively metabolizing and actually exist
in a state of dormancy. Ecosystems experience states of nutrient flux, and starvation due to
nutrient limitation is a cue that regulates dormancy in natural communities (Braeken et al. 2006).
Theory predicts benefits afforded by dormancy should be greater in environments with limited or
low resource availability (Gardner et al. 2007). Soil environments can be especially
heterogeneous, with changes in soil properties affecting water flow and nutrient cycling
(Schlesinger et al. 1996). Metagenomic analysis of sequencing data from different environments
114
revealed a high abundance of dormancy genes present in soil environments (Lennon & Jones
2011). An effective life detection strategy must not only be able to detect metabolically active
life, but potentially starving or dormant life as well.

With this study, we investigated how the addition of an electron donor would potentially
stimulate microbial activity of dormant populations and consequently be detected by a MFC
system. Non-sterile fuel cell experiments where no lactate (Figure 4.3) is added look very similar
to sterile MSS soil experiments with lactate addition, indicating that redox reactions between the
soil and organics can provide false positive results (Figure 4.5B). The sterile HF soil with lactate
addition did not show the same response as the sterile MSS soil, suggesting that perhaps the
perchlorate present in the MSS salt solution may react with some of the lactate causing the
gradual increase in current density. When lactate is added to the non-sterile fuel cells, a large and
unmistakable response occurs, demonstrating the usefulness of supplying an additional electron
donor, and not just relying on what is in the soil itself (Figures 4.4D and 4.5D). The enhanced
current density (four times the current observed in long term tests) in both non-sterile soils
highlights the reproducibility of microbial activity profiles, with increased current that is
maintained over time. The use of lactate for metabolic activity is likely when compared to the
controls; adding a buffer (PIPES) did not change current density signal in non-sterile soils and
with the sterile soils no sustained current density is observed at the level of the non-sterile
experiments with the addition of lactate. HPLC results also revealed an accumulation of lactate
in the sterile soils compared to the non-sterile soils. Pyruvate and acetate were both measured in
the non-sterile soil lactate addition experiments, and both organic acids are the byproducts of
lactate oxidation. Their presence indicates the potential biotic oxidation of lactate. These
metabolic byproducts of lactate oxidation could also be used by other organisms in the microbial
115
community, stimulating further metabolic activity.

It is important to note that to be able to distinguish between abiotic and potentially biotic signals,
BECD experiments should run on longer timescales. If these experiments were only run for a
couple of hours, it would be easy to misinterpret some of the current density observed. In
addition, the current density from abiotic reactions usually occurred in large spikes that decay
rapidly, while current density from metabolic activity exhibited a sustained response indicative
of cellular activity and growth. These abiotic spikes often occurred when there was an increased
rate of mixing by injecting test substrates into the anode chamber. The only consistent mixing in
the MFC systems was contributed by the purging with N
2
gas. Evaporation of the anolytes could
also disrupt the electrical signals detected as seen towards the end of the time course in Figure
4.5B. It is important to note that determining the appropriate controls for these experiments is
complicated by the fact that standard sterilization through autoclaving seemed to change the
electrochemical baseline of the soils (Figure 4.2). CV tests on the soil substrates showed that
oxidants were released upon sterilization of the soils, which can further complicate interpretation
of the electrochemical signals observed in full cell tests. The organic acid addition tests
conducted with an organic acid and a control buffer, demonstrated that this experimental design
might be sufficient to distinguish between abiotic and biotic electrochemical signals. For future
instrumentation, implementing these types of controls rather than heat sterilization may help
simplify and reduce the overall required instrument payload.

Quantification of the proteins present in the soil anolyte solution and on the electrodes allowed
us to approximate changes in biomass. Overall soil anolyte protein concentrations were higher in
the non-sterile soil with lactate addition experiments. Non-sterile MSS soil with lactate anolyte
116
protein concentrations were much higher than the concentrations for other conditions at all time
points. Accordingly, the current density observed for the non-sterile MSS soil lactate addition
experiments was also the highest achieved throughout the time course of the MFC experiments.
Electrode protein concentrations were highest for both soils under non-sterile conditions with
lactate addition, although concentrations were also high with the lactate additions to sterile soil
experiments. The lack of biofilms evident in SEM images of the electrodes, along with the
anolyte protein data, suggest that the planktonic community might have contributed the most to
current density. Many soils are rich in humic acids that can serve as electron shuttles (Klüpfel et
al. 2014). Electron shuttles such as humic acids are known to mediate electron transfer between
microbes and insoluble electron acceptors/donors (Kappler et al. 2004, Wolf et al. 2009). We
also explored introducing exogenous electron shuttles (riboflavin and flavin mononucleotide)
into our MFC systems but no enhancement in current density was observed (data not shown).

Experiments using different cell abundances of S. oneidensis MR-1 were conducted to determine
the sensitivity of BECD systems to cell number. Estimates of cell numbers in marine subsurface
sediment suggest that cell numbers drop according to the power law, with 10
9
cells/cm
3
near the
sediment surface to 10
6
cells/cm
3
hundreds of meters below (Whitman et al. 1998; Parkes et al.
2000). In the South Pacific Gyre, one of the most oligotrophic regions on Earth, the number of
cells found in sediments 1 m below the seafloor is estimated to be between 10
6
-10
3
cells/mL
(Kallmeyer et al. 2012). Multiple starting cell inoculums were utilized in our experiments from a
range of 10
5
to 10
6
cells/mL. Higher cell numbers provided greater signal initially, but with time
the lower cell inoculums generated greater current density (Figure 4.8). These results might be
due to resource depletion, as higher cell numbers can initially quickly consume the electron
donor provided. Lower cell numbers might be slower to multiply and oxidize the electron donor
117
available, but can take advantage of the resources later showing a delayed signal in current. For
the detection of life purposes, resources may be more constraining than cell number. Future
experiments are needed to track cell number during the time course of the experiments to better
understand how current density may change with cell number, and step down starting cell
numbers to even lower numbers (< 10
5
cells/ml).

CONCLUSION

BECDs are a promising approach to life detection that can be independent of the metabolic
modes present in an environment, and complementary to biochemical approaches. In concert
with habitability studies, BECD experiments can also be designed to probe for specific
metabolisms. Habitability studies might indicate what dominant electron donors or acceptors are
present in the environment, but typically if life is using a particular substrate in its metabolism,
concentrations should be changing and/or these compounds may elude detection. For example, a
site might have ample electron acceptors but no measurable electron donors. By determining
what might be a likely or favorable electron donor in an environment, addition tests similar to
our lactate experiments can be conducted to stimulate metabolic activity. In addition, BECDs
like a three-electrode chemical cell can potentially test various redox potentials (electron donors
or acceptors) in an environment in situ. If a particular redox potential can be used for energy by
life, current should be easily detected and measurable.

118
APPENDICES

Appendix S1. Additional baseline full cell tests conducted on: A) & B) Sterile and Non-
Sterile HF Soil and C) & D) Sterile and Non-Sterile MSS.
119
Appendix S2. Representative SEM images of the carbon felt electrodes after long-term
microbial full cell experiments. A) & B) Non-sterile HF full cell carbon felt electrode, C) & D)
Sterile HF full cell carbon felt electrode. Cell like structures are circled in yellow.

A)
B)
C)
D)
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CHAPTER 5

CONCLUSION AND FUTURE PERSPECTIVES

The work presented in this thesis demonstrates the value of using electrochemical techniques to
investigate microbial metabolism, more specifically the process of extracellular electron
transport (EET) and insoluble substrate oxidation. Although EET has been characterized and
described in metal reducing microorganisms, current evidence suggests the process of EET is
widespread in nature and present across phylogenies (Gralnick & Newman 2007). EET might be
especially important in environments where organic matter is scarce. For example, due to the
limiting organic electron donors available in the most deep-sea sediments, solid phase mineral
electron donors have been suggested to be important electron sources (Orcutt et al. 2011; Orcutt,
LaRowe, et al. 2013; Orcutt, Wheat, et al. 2013). However, the lack of knowledge surrounding
the process of insoluble substrate oxidation, coupled with the difficulty in monitoring redox
changes in solid substrates, inhibits our knowledge of the importance and impact of EET on
various ecosystems.

By combining electrochemistry with microbiological and molecular methods, the studies
presented in this thesis have expanded our understanding of what microorganisms are capable of
insoluble substrate oxidation. Utilization of these techniques, led to the successful enrichment
and isolation of six novel strains capable of insoluble substrate oxidation from one type of
marine sediment. Genera such as the Vallitalea have not been recognized as EET-capable, and
other genera such as the Arcobacter and Vibrio have few reports of EET-activity with those all
being related to insoluble substrate reduction (Fedorovich et al. 2009; Wee et al. 2014). By
identifying and isolating these new strains, we can start to probe environmental 16S rRNA and
metagenomic datasets for the presence of these OTUs and assess if they might be important
125
players in these systems. Additionally, two more Marinobacter sp. were isolated, adding to the
repository of Marinobacter capable of cathode oxidation (Rowe et al. 2015). Whole genome
sequencing of these strains has begun and will provide a starting point to identifying key genes
and pathways involved in insoluble substrate oxidation. These Marinobacter strains are
facultatively aerobic heterotrophs, making them amenable and ideal candidates for genetic work
and mutation-based studies to fully elucidate new EET mechanisms.

The three-electrode system enrichment technique utilized in this work can easily be applied to
other environmental samples of interest (e.g. sediments beneath low productivity gyres) or even
be conducted in-situ. This approach can facilitate investigation of microbes that are often elusive
to culturing. The application of multiple biological redox potentials of interest was instrumental
in our work to expand and more fully capture the diversity of cathode-oxidizing microorganisms.
In addition, a culture-independent approach combining metagenomics and metatranscriptomics is
being implemented to study cathode-oxidizing communities from a deep-sea, whale fall
sediment. The application of these molecular techniques to electrode communities will help
identify transcriptionally active putative mineral-oxidizing microbes and gene targets that are
differentially regulated in response to EET-stimuli. It is our hope that the work presented in this
thesis will continue to be built upon and expanded. The growth of the field of
electromicrobiology is critical for understanding the diversity of EET-capable microbes and EET
processes in the environment. The knowledge gained will also potentially help optimize the use
of these microbes and bioelectrochemical systems to tackle problems such as pollution,
wastewater treatment, and the development of sustainable alternative energy.

126
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Abstract (if available)

Abstract The influence of microorganisms on our world is tremendous as they are primary drivers of biogeochemical cycles and regulators of ecosystem processes. Their influence comes from the diverse and often complex metabolic capabilities that they possess, allowing the utilization of a wide range of substrates for the derivation of energy. The substrates they can use as either electron donors or acceptors was once thought to be limited to those that were soluble and could readily enter the cell membrane. The discovery of extracellular electron transport (EET) has broadened the diversity of substrates to include insoluble substrates such as minerals and metals. EET is the ability of microbes to transfer electrons to and from insoluble substrates outside of the cell. Much of the knowledge about EET has been gained through investigations of model organisms from two genera, Geobacter and Shewanella, and these studies have mainly focused on how electrons are transferred from cells to insoluble substrates. The process of receiving electrons from insoluble substrates has largely remained unexplored, and the mechanisms of solid substrate oxidation are poorly characterized and understood. ❧ The goal of this dissertation work was to further understanding of EET with a focus on cathode (insoluble substrate) oxidation. Microbes capable of insoluble substrate oxidation have been elusive to culturing, in part due to the difficulty in maintaining the conditions they require in the laboratory. Electrochemical techniques have allowed us to mimic the redox conditions provided by solid substrates and study the process of insoluble substrate oxidation in a quantifiable and controllable manner. Combining electrochemical techniques with microbiological and molecular methods, I was able to investigate the influence of redox potential on cathode-oxidizing community structure. Overall community diversity and richness increased with more negative applied redox potentials. In addition, abundances of important known EET groups, including the Altermonadales, Clostridiales, and Desulfuromonadales, varied with redox potential. Motility and chemotaxis genes were found in greater abundance in electrode communities, suggesting the importance of these pathways for colonization and utilization of the electrode as an electron donor. The cathode-oxidizing community enrichments demonstrated the validity of this approach in capturing groups that are known to participate in EET and also highlighting potentially important novel groups (e.g. Campylobacterales) that perform EET as well. The initial enrichments and the insights gleaned from molecular analyses laid the foundation for the successful isolation of six new strains of bacteria from five different classes capable of cathode oxidation. These isolates are not only phylogenetically diverse, but they also display varying electrochemical properties suggesting different mechanisms of EET. The microorganisms highlighted in this work will help inform potential genetic markers for future studies as well as aid in developing a framework for detecting EET capabilities in environmentally relevant microbes. The potential application of bioelectrochemical devices (BECDs) to probe for microbial metabolic activity in environmental samples was also explored. The proof of concept investigation demonstrated the utility of BECDs as a life detection strategy. This work contributes to the modernization and improvement of life detection techniques focused on metabolism. The data herein presented reveals the importance of EET as a widespread metabolic ability and environmental process, which is only starting to be understood with the advances made in electromicrobiology.

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Asset Metadata

Creator Lam, Bonita Rasmey (author)

Core Title Extracellular electron transport: Investigating the diversity and mechanisms behind an understudied microbial process with global implications

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Biology

Publication Date 01/18/2018

Defense Date 12/08/2017

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

Tag electromicrobiology,environmental microbiology,extracellular electron transport,marine sediment,microbial metabolism,OAI-PMH Harvest

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Nealson, Kenneth (committee chair)

Creator Email bonitarl@usc.edu,bonitarlam@gmail.com

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c40-463318

Unique identifier UC11266821

Identifier etd-LamBonitaR-5957.pdf (filename),usctheses-c40-463318 (legacy record id)

Legacy Identifier etd-LamBonitaR-5957.pdf

Dmrecord 463318

Document Type Dissertation

Rights Lam, Bonita Rasmey

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