BIOLOGICAL MECHANISMS OF URANIUM TRANSFORMATION CATALYZED BY
GEOBACTER BACTERIA
By
Dena L. Cologgi

A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
DOCTOR OF PHILOSOPHY
Microbiology and Molecular Genetics
2012

ABSTRACT
BIOLOGICAL MECHANISMS OF URANIUM TRANSFORMATION CATALYZED BY GEOBACTER
BACTERIA
By
Dena L. Cologgi
An insufficient knowledge of the biological mechanisms of contaminant transformation
often limits the performance of in situ subsurface bioremediation and long-term stewardship
strategies. The in situ stimulation of Fe(III) oxide reduction by Geobacter bacteria leads to the
concomitant precipitation of U(VI) from groundwater. However, the biological mechanism
behind this reaction has remained elusive. Because Fe(III) oxide reduction requires the
expression of conductive pili in Geobacter, we also evaluated their contribution to uranium
reduction. In chapter 2 of my dissertation I demonstrate a previously unrecognized role for
Geobacter pili in the extracellular reduction of uranium and its essential function as a catalytic
and protective cellular mechanism.
The expression of pili by Geobacter also promotes cell aggregation and biofilm
formation. Recent work has shown that Geobacter cells transition from planktonic to biofilm
physiologies during the active phase of U reduction in the subsurface. Despite these findings,
the contribution of Geobacter biofilms to uranium removal and reduction has not been
investigated. In chapter 3 of my dissertation I demonstrate that multilayer biofilms are able to
reduce and tolerate substantially more U than planktonic cells for prolonged periods of time,
making them an attractive option for the development of permeable biobarriers for U
bioremediation. I also demonstrate the role of pili as a primary U reductase in the biofilm.

To gain further insight into how biofilms transform U, in chapter 4 of my dissertation I
screened a library of transposon-insertion mutants and identified mutants with biofilm defects.
This study confirmed the role of Geobacter pili in biofilm formation, and identified other genes
encoding cell envelope and electron transport components that had not previously been
implicated in biofilm development. These molecular markers can be used to predict and
monitor the physiological state of Geobacter bacteria during the in situ bioremediation of U.
Previous work, including the prior chapters of my dissertation, has highlighted the
importance of the cell envelope and its components for the survival of Geobacter in the
subsurface. However, little is known regarding the regulation of the cell envelope. Thus, I
investigated the role of the Geobacter’s ECF sigma factor, RpoE. In the last chapter of my
dissertation, I show that RpoE is required for response to cell envelope stress, as well as the
regulation of Geobacter’s extracellular electron transfer pathways. This highlights the functional
specialization that RpoE has undergone to control the adaptive responses that enable
Geobacter bacteria to survive in the environment, and links my findings to the physiology of
Geobacter in the subsurface.

DEDICATION

This dissertation is dedicated to my family. Thank you for staying rational when I wasn’t, and
believing in me when I didn’t. I am so fortunate to have you in so many ways. I am incredibly
thankful that being in Michigan has given me the opportunity to know my family in New York
and become a part of their lives, especially during this past year. Thank you for all of your love,
guidance and support.

iv

ACKNOWLEDGEMENTS

I would first like to acknowledge my mentor, Dr. Gemma Reguera, who has been there
for me throughout this process and taught me more than I can describe. Thank you for taking a
tremendous risk and entrusting your first lab to two very enthusiastic (but even more
inexperienced and naive!) first-year rotation students. I am proud to have been a part of that
beginning. I also need to extend a huge thank you to Allison Speers who has been with me since
day one when we started the transposon mutagenesis together during our rotation. I don’t
think I would have survived grad school without you, Allison – literally! I will always owe you for
accompanying me to Argonne and ensuring I didn’t do something horrible (hitting the big red
button, locking myself in the hutch, blowing up the synchrotron...) during those late-night
sample changes performed in a nearly non-functional, sleep-deprived state.
I would also like to thank the other individuals who contributed directly to the work
presented in my dissertation. Blair Bullard worked side-by-side with me on many of the biofilm
experiments, and was instrumental in obtaining anything and everything needed for my
research! Two amazing undergrad students, Annie Otwell and John Rotondo worked with me
on the transposon mutagenesis project. Annie and John, you made my first experiences
mentoring both easy and fun; I was very lucky to have you. No one else in the world would have
so calmly tolerated my ups and downs, or endured so well the endless colony picking, plate
transfers, and rarely-successful sequencing attempts. I would also like to acknowledge Sanela
Lampa-Pastirk, who developed the pili isolation protocol used in chapter 2. Sanela welcomed

v

me into her family and provided significant support for me, both emotionally as well as
scientifically, throughout this process.
I would like to thank our collaborator, Shelly Kelly (EXAFS analysis) not only for her
scientific contributions, but also for taking the time to train (and in general put up with) Allison
and me so we could play a more active role in our trips to Argonne. And special thanks for being
such a wonderful hostess and humoring us in our non-scientific adventures including the search
for albino deer, the tricycle rides, and the hunt for the coffee cart! I’d also like to acknowledge
the rest of the Sector 20 staff at Argonne National Lab for all of their help and patience
Acknowledgement must also go to the other members of the Reguera Lab who may not
have been directly involved in my research, but have had a huge impact nonetheless. First, to
my long-term lab-mates Jenna Young, Becky Steidl, and Mike Manzella, along with Allison and
Sanela -- what can I say? I can’t imagine a lab without you. You have truly been like family.
Thank you so much for your humor, support and loyalty. I would never have survived without
you. I would also like to thank Bryan Schindler, Jihwan Hwang, Ana Lara, and Catherine Silva.
Though they were in our lab only a short time, I am very thankful for both their scientific input
and conversations about life. I have to especially thank Ana for keeping me company during
those depressing late night battles with the KPA! I would also like to acknowledge our
dishwashers and media-makers (Marvin, Max, Derek, Mike2, Mellissa, Karl, Katie, Alex and Eric).
Without them our lab couldn’t function. The last in this category is Kwi Kim, who has
relentlessly kept me on track and made sure that I didn’t forget what’s important in life.
Other people who have been integral to the success of my grad school career include
my committee members, Dr. Rob Britton, Dr. Terry Marsh, and Dr. Lee Kroos, who have been
vi

very patient and supportive throughout this process. I would also like to acknowledge several
individuals who assisted with some of the technical aspects of my research including Matthew
Marshall (PNNL), Evgenya Shelobolina (University of Wisconsin-Madison) and Kazem Kashefi,
who provided some of the protocols crucial to my work, as well as Weimin Chen (and Sanela
and Igor) who helped me master the KPA. I would also like to extend a huge thank you to the
amazing people at the Center for Advanced Microscopy (Alicia Pastor, XouDong Fan, Melinda
Frame, Carole and Stanley Flegler) who have gone out of their way to be helpful, and were
always understanding when I was running very late! As Alicia (who I also need to thank for
being such a great friend and neighbor!) once so eloquently put it, “this is the Center for
Advanced Microscopy, where all your dreams come true!”
I have been fortunate enough to have been funded throughout my Ph.D. by agencies
such as the National Institute of Environmental Health Science’s Superfund program (R01
ES017052-03), the Office of Science (BER), U.S. Department of Energy (DOE) (DE-SC0000795),
the College of Natural Science at Michigan State University (Hensley fellowship), and the
Biogeochemistry Environmental Research Initiative at Michigan State University (research
fellowship). PNC/XSD facilities and research at the APS are supported by the US DOE-BES, a
Major Resources Support grant from NSERC, the University of Washington, Simon Fraser
University and the APS. The APS is an Office of Science User Facility operated for the U.S. DOE’s
Office of Science by Argonne National Laboratory and supported by the U.S. DOE under
contract No. DE-AC02-06CH11357.

vii

I, of course, also have to acknowledge the other MMG grad students who have been
with me over the years. I am incredibly grateful for your camaraderie, companionship, support
and scientific advice, and I wish you all the best.
Lastly, I would like to thank my family, to whom this dissertation is dedicated.

viii

TABLE OF CONTENTS
LIST OF TABLES ..........................................................................................................xii
LIST OF FIGURES ....................................................................................................... xiii
KEY TO SYMBOLS AND ABBREVIATIONS .................................................................... xvi
CHAPTER 1
INTRODUCTION .......................................................................................................... 1
URANIUM CONTAMINATION AND ITS IMPACT ON PUBLIC HEALTH .................. 2
REMEDIATION STRATEGIES.............................................................................. 3
MICROBIAL U(VI) REDUCTION.......................................................................... 4
THE SEARCH FOR U(VI) REDUCTASES................................................................ 6
PILUS NANOWIRES .......................................................................................... 7
BIOFILMS IN THE SUBSURFACE ........................................................................ 9
BIOFILM DEVELOPMENT ................................................................................ 11
REGULATION OF CELL ENVELOPE COMPONENTS AND ELECTRON TRANSFER
PROCESSES .................................................................................................... 12
REFERENCES .................................................................................................. 13
CHAPTER 2
EXTRACELLULAR REDUCTION OF URANIUM VIA GEOBACTER CONDUCTIVE PILI AS A PROTECTIVE
CELLULAR MECHANISM ............................................................................................... 21

ABSTRACT ..................................................................................................... 22
INTRODUCTION ............................................................................................. 23
MATERIALS AND METHODS ........................................................................... 25
Strains and culture conditions .................................................................. 25
Pili induction at suboptimal growth temperatures .................................. 26
U(VI) resting cell suspension assays ......................................................... 26
Gene expression analyses by qRT-PCR ..................................................... 27
TEM and Energy Dispersive Spectroscopy (EDS) analyses........................ 28
Pili purification, quantification, and biochemical characterization .......... 28
Transmission Electron Microscopy (TEM) and Confocal Laser Scanning
Microscopy (CLSM) analyses of purified pili ............................................. 31
SDS-PAGE and staining of outer membrane, heme-containing proteins . 31
X-ray Adsorption Spectroscopy (XAS) analyses ........................................ 32
Vitality and viability assays ....................................................................... 33
RESULTS ........................................................................................................ 34
Expression of pili promotes the extracellular reduction of U(VI) ............. 34
X-ray absorption fine structure (EXAFS) analyses demonstrate the
reduction of U(VI) to mononuclear U(IV) ................................................. 41
U reduction via pili as a cellular protective mechanism ........................... 45
ix

DISCUSSION .................................................................................................. 47
Physiological relevance of the extracellular reduction of U by Geobacter’s
pili .............................................................................................................. 47
Reduction of U to mononuclear U(IV) phases .......................................... 48
Model for the reduction of U(VI) by Geobacter bacteria ......................... 50
Implications for the in situ bioremediation of U ...................................... 52
REFERENCES .................................................................................................. 53
CHAPTER 3
URANIUM TRANSFORMATIONS DURING BIOFILM DEVELOPMENT IN GEOBACTER
SULFURREDUCENS .................................................................................................... 61
ABSTRACT ..................................................................................................... 62
INTRODUCTION ............................................................................................. 63
MATERIALS AND METHODS ........................................................................... 64
Strains and culture conditions .................................................................... 64
U(VI) resting cell suspension and biofilm assays ......................................... 66
Vitality fluorescent assays .......................................................................... 66
Microscopy ................................................................................................ 67
X-ray Absorption Spectroscopy (XAS) analyses ........................................... 68
SDS-PAGE and heme-stain of proteins of the biofilm EPS matrix ................. 69
RESULTS AND DISCUSSION ............................................................................ 70
Enhanced U(VI) immobilization and tolerance by biofilms .......................... 70
Extracellular reduction of U by biofilms ...................................................... 71
Role of conductive pili and c-cytochromes of the biofilm matrix in U
reduction ................................................................................................... 76
XAS analyses .............................................................................................. 78
Implications for bioremediation ................................................................. 81
APPENDIX ..................................................................................................... 82
REFERENCES .................................................................................................. 88
CHAPTER 4
GENETIC ANALYSIS OF BIOFILM FORMATION IN GEOBACTER SULFURREDUCENS ....... 93
ABSTRACT ..................................................................................................... 94
INTRODUCTION ............................................................................................. 95
MATERIALS AND METHODS ........................................................................... 98
Strains and culture conditions .................................................................. 98
Biofilm assays ............................................................................................ 98
Confocal Laser Scanning Microscopy (CLSM) ........................................... 99
Transposon mutagenesis .......................................................................... 99
High-throughput biofilm screening ........................................................ 100
Identification of transposon-insertion sites ........................................... 101
Rescue cloning ............................................................................ 101
Arbitrary PCR ............................................................................... 102
Mutant confirmation .............................................................................. 103
x

SDS-PAGE and heme-stain of proteins in biofilm EPS matrix ................. 104
U(VI) resting cell suspension biofilm assays ........................................... 104
RESULTS AND DISCUSSION .......................................................................... 105
Development of an assay to characterize biofilm developmental
stages ...................................................................................................... 105
Identification of Tn5-insertion mutants with biofilm defects ................ 108
Description of mutant groups ................................................................. 109
Cell envelope............................................................................... 110
Electron transfer ......................................................................... 111
Role of pili in biofilm development and uranium immobilization .......... 114
Conclusions ............................................................................................. 117
APPENDIX ................................................................................................... 118
REFERENCES ................................................................................................ 127
CHAPTER 5
IDENTIFICATION AND CHARACTERIZATION OF THE RPOE SIGMA FACTOR OF GEOBACTER
SULFURREDUCENS .................................................................................................. 134
ABSTRACT ................................................................................................... 135
INTRODUCTION ........................................................................................... 136
MATERIALS AND METHODS ......................................................................... 138
Bacterial strains and culture conditions ................................................. 138
Complementation ................................................................................... 138
Transcriptional analysis of the rpoE gene and the gene region by
RT-PCR ..................................................................................................... 139
Stress tolerance assays ........................................................................... 140
Biofilm assays .......................................................................................... 141
Transmission Electron Microscopy (TEM) .............................................. 141
Fe(III) reduction assays ........................................................................... 142
RESULTS AND DISCUSSION .......................................................................... 142
Genetic organization of the rpoE region................................................. 142
RpoE is not essential for growth, but regulates cell envelope integrity
during growth transition ......................................................................... 144
RpoE is required for stress tolerance...................................................... 149
RpoE regulates electron transfer at the cell envelope ........................... 154
Implications ............................................................................................. 156
REFERENCES ................................................................................................ 158
CHAPTER 6
CONCLUSIONS AND FUTURE DIRECTIONS................................................................ 166
REFERENCES ................................................................................................ 171

xi

LIST OF TABLES
2

*........................................................ 44

TABLE 2.1

EXAFS modeling results for R and σ

TABLE 2.2

EXAFS modeling results for coordination numbers .................................... 44

TABLE 3.1

COMSTAT analysis ....................................................................................... 75

TABLE 3.A.1 EXAFS modeling results for R and σ

2

** ..................................................... 87

TABLE 3.A.2 EXAFS modeling results for coordination numbers .................................... 87
TABLE 4.1

Primers ...................................................................................................... 102

TABLE 4.A.1 Plates, coatings, growth media, and solvents assessed for their use in the
biofilm assay .............................................................................................. 119
TABLE 4.A.2 Transposon-insertion mutants displaying deficient or enhanced biofilm
phenotypes................................................................................................ 122

xii

LIST OF FIGURES
FIGURE 2.1

Reduction and subcellular localization of U .............................................. 35

FIGURE 2.2

Fold change expression of pilA relative to recA in resting cell suspensions
of the WTP+ and WTP- strains after 6 h of incubation with 1 mM U(VI)
acetate ....................................................................................................... 36

FIGURE 2.3

Micrographs of purified pilus fibers .......................................................... 37

FIGURE 2.4

Energy dispersive X-ray spectrum of the pili-associated electron-dense
deposits imaged by TEM .......................................................................... 38

FIGURE 2.5

Subcellular localization of uranium deposits by TEM analyses of thin
sections of pili-expressing (WTP+ and pRG5::pilA) and non-expressing
-

(WTP- and PilA ) cells ................................................................................. 39
FIGURE 2.6

SDS-PAGE profiling of mechanically detached outer membrane
proteins ..................................................................................................... 40

FIGURE 2.7

U LIII-edge EXAFS spectra and models ...................................................... 42

FIGURE 2.8

Effect of U(VI) exposure on cell vitality and viability ................................ 46

FIGURE 2.9

Model for the extracellular reduction of U(VI) to U(IV) ............................ 50

FIGURE 3.1

Biofilms maintain vital activities through 24 h of U exposure .................. 70

FIGURE 3.2

SEM micrographs of 48 h biofilms ............................................................ 72

FIGURE 3.3

U(VI) immobilization/reduction, and cytochrome profile of the WT, PilA , and
pRG5::pilA strains ...................................................................................... 73

FIGURE 3.4

CLSM micrographs showing side view projections of (A) WT (24, 48 and 72h),

-

-

(B) PilA (48 h), and (C) pRG5::pilA (48 h) biofilms stained with the BacLight
viability kit (green, live cells; red, dead cells) ............................................ 74
FIGURE 3.5

U LIII-edge XAFS spectra. EXAFS spectra and model .................................. 79

FIGURE 3.A.1 SEM micrograph and EDS analysis of U precipitates ................................. 83
xiii

FIGURE 3.A.2 CLSM micrographs showing top view projections of 24, 48, and 72 h WT (A),
-

48 h PilA (B) and 48 h pRG5::pilA biofilms stained with BacLight viability kit
(green, live cells; red, dead cells) ............................................................ 84
FIGURE 3.A.3 Correlation of biofilm age and biofilm biomass (A), thickness (B), surface
area (C), surface coverage (D), and surface:volume ratio (E) ................. 85
FIGURE 3.A.4 Linear correlation between the total protein content of biofilms and
2

biofilm age (R =0.94) (A) and between piliation (planktonic cells grown
under pili-inducing conditions at 25°C) and U reduction by 48 h biofilms
-

2

of the WT, PilA and pRG5::pilA strains (R = 0.999) (B) ......................... 86
FIGURE 4.1

Comparison of the effect of growth medium type and electron donor
(acetate) concentrations on biofilm formation (A). CLSM images of
wild-type biofilms grown in FW 30A/40F (B) (24 (top), 48 (middle), and
72 (bottom) h growth), FW 15A/40F (72 h growth) (C), and DB 30A/40F
(72 h growth) (D). Bar, 20 µM ................................................................ 107

FIGURE 4.2

Crystal violet staining of representative Tn5-insertion mutants ........... 109

FIGURE 4.3

Phenotypic characterization of selected mutants ................................. 116

FIGURE 4.A.1 A selection of 96-well plates investigated for their ability to maximize
G. sulfurreducens 72-hour biofilm growth at 30°C ................................ 120
FIGURE 4.A.2 Selection of a solvent for de-staining of crystal violet biofilms ............. 121
FIGURE 4.A.3 Southern blot to confirm a single Tn5 insertion event .......................... 126
FIGURE 5.1:

Genetic arrangement of the rpoE region ................................................ 142

FIGURE 5.2:

RpoE demonstrates impaired growth in NBAF medium compared to
WT........................................................................................................... 145

FIGURE 5.3:

Regulation of biofilm formation by RpoE ............................................... 148

FIGURE 5.4:

The effect of environmental stressors on the growth of the wild-type
(solid symbols) and the RpoE- mutant (open symbols) strains .............. 151

FIGURE 5.5:

Reverse transcriptase (RT)-PCR of rpoE and recA transcripts................. 153

-

xiv

FIGURE 5.6:

Fe(III) reductase activity of resting cell suspensions............................. 154

xv

KEY TO SYMBOLS AND ABBREVIATIONS
CLSM…………………………………………………………………………………. Confocal Laser Scanning Microscopy
DB (30A/40F) ……………………….......... minimal medium with 30 mM acetate and 40 mM fumarate
EPS………………………………………………………………………………………………………………… exopolysaccharide
EXAFS………………………………………………………………………. Extended X-Ray Absorption Fine Structure
Fe(II), Fe(III)…………………………………………………. iron in the +2 and +3 oxidation states, respectively
FWAF or FW (15A/40F) …………… fresh water medium with 15 mM acetate and 40 mM fumarate
FW (30A/40F) ……………………….... fresh water medium with 30 mM acetate and 40 mM fumarate
L…………………………………………………………………………………………………………………………………………… liter
ml…………………………………………………………………………………………………………………………………. milliliter
mM…………………………………………………………………………………………………………………………….. millimolar
mol…………………………………………………………………………………………………………………………………….moles
NBAF…………………………………………………. nutrient broth with 15 mM acetate and 40 mM fumarate
nm……………………………………………………………………………………………………………………………. nanometer
OD580………………………………………………………………………………………………… optical density at 580nm
OD600………………………………………………………………………………………………… optical density at 600nm
PCR…………………………………………………………………………………………………… polymerase chain reaction
-

PilA …………………………………………………………………………………………………………. pilin-deficient mutant
SEM…………………………………………………………………………………………….. Scanning Electron Microscopy
TEM………………………………………………………………………………………. Transmission Electron Microscopy
U…………………………………………………………………………………………………………………………………… uranium
xvi

U(IV), U(VI)……………. uranium in the +4 (insoluble) and +6 (soluble) oxidation state, respectively
µg……………………………………………………………………………………………………………………………… microgram
µl………………………………………………………………………………………………………………………………… microliter
XANES………………………………………………………………………………. X-ray absorption near edge structure

xvii

CHAPTER 1

INTRODUCTION

1

URANIUM CONTAMINATION AND ITS IMPACT ON PUBLIC HEALTH
Uranium (U) is a naturally occurring metal that is nearly ubiquitous at low levels in soil
and rocks (1). U is an extremely dense metal, which serves as an excellent large-scale and
emission-free power source (25). Thus, it is frequently used for both civilian (nuclear reactors)
and military (nuclear submarine reactor cores, nuclear weapons, radiation shielding, missile
projectiles) applications (1, 34). From the 1940s on, particularly during the Cold War era, U has
been extensively mined for its use in the production of the items listed above, leaving behind
large amounts of radionuclide waste as a result. Other sources of nuclear waste and
contamination include weapons testing, fuel reprocessing, as well as nuclear accidents such as
Chernobyl (25). The largest volume of waste, however, is the result of U mining and milling
processes (25).
Although many believe the primary health risk from U lies in its radioactivity, in general
it is the chemical toxicity that is more hazardous (34). Toxicity can occur via inhalation,
ingestion, or through direct contact with the skin. Following a significant acute or chronic
exposure event, serious health effects can occur. The most commonly affected organs are the
kidneys, but in some cases the respiratory, neurological, and reproductive system can also be
affected (1). For this reason, the U.S. Environmental Protection Agency (EPA) has established a
maximum allowable level of 30 µg/L in drinking water (1), and the Superfund program has set a
preliminary remediation goal of 2.22 µg/L (34).

2

REMEDIATION STRATEGIES
The mobility of U in the subsurface is dependent on a variety of factors including pH,
redox potential, and other materials present (coatings, sediments, organic and inorganic
compounds) (1, 34, 69). Under oxidizing conditions (i.e. surface or groundwater) U is present in
2+

its soluble form as a uranyl ion (U(VI) or UO2 ) or in carbonate complexes. When reduced by
biotic or abiotic factors U is instead present in the insoluble uranous (U(IV)) state (34, 69). It is
soluble U that poses a greater public health threat as it can migrate in groundwater and
contaminate larger areas, as well as drinking water supplies. For this reason, many
bioremediation schemes seek to reduce U(VI) to U(IV), thereby immobilizing it and preventing
its migration.
Since radionuclides such as U cannot be degraded, and have a lengthy half-life, removal
or sequestration of the contaminant is the only available remediation strategy. This can be
accomplished using physical or chemical processes (i.e. containment, excavation, chemical
extraction, chelation), although these methods are often costly, non-specific, and can produce
large amounts of waste which require safe disposal (4, 19, 37, 69). Biological methods,
however, are often a safer, more cost-effective option for either in situ or ex situ remediation,
and in some cases can be used as a complement to physical and chemical methods (19). In situ
remediation is especially advantageous as it requires little excavation of the affected site, and a
smaller risk of exposing the public and/or remediation team to the contaminant (19). Biological
mechanisms of bioremediation include phytoremediation (the sequestration of the
contaminant by plants), or the stimulation of natural microbial communities to convert the
contaminant either to a less toxic, or less soluble form. As described below, microbial
3

bioremediation is a promising strategy. However, a lack of knowledge regarding the mechanism
of U reduction by subsurface bacteria has impaired the success of these techniques.

MICROBIAL U(VI) REDUCTION
Although the reduction of U(VI) in anaerobic environments has long been recognized, it
was initially presumed that the observed U reduction was due to an indirect and abiotic
association with the microorganisms (29). It was believed that microbes acted as passive
attachment surfaces for abiotic reduction, or provided reduced metabolic byproducts (such as
H2S) that in turn were responsible for the observed U reduction (29, 38). This premise was later
challenged with the discovery that dissimilatory metal reducers such as GS-15 (later renamed
Geobacter metallireducens) and Alteromonas putrefaciens (later renamed Shewanella
putrefaciens) were capable of using U as an energy source, and could carry out U reduction
more quickly than was observed for abiotic processes. This demonstration of the direct role of
bacteria in U reduction led the researchers to hypothesize that this phenomenon could be
harnessed for bioremediation purposes (38).
Following these investigations, experiments were carried out under more
environmentally relevant conditions, specifically within sediment microcosms in the laboratory.
These studies demonstrated that microbial reduction of U followed the consumption of
acetate, but also noted the concomitant reduction of Fe(III) (17, 18). This lends support to the
idea that U reduction is carried out by dissimilatory metal-reducing bacteria, and it was
hypothesized that the Fe(III) in the environment was providing the energy for bacterial growth
during in situ stimulation. This is likely because Fe(III) was present in significantly higher
4

quantities (mmol/kg sediment) than the U (µmol/kg sediment) (17). Consistent with the
observed Fe(III) and U(VI) reduction, 16S ribosomal DNA sequences obtained from acetateamended sediments demonstrated an increase (40% of sequences compared to 5% in
untreated sediments) in the Geobacteracea, a family which includes dissimilatory metalreducers such as the organisms described above (17, 18). These laboratory-scale microcosm
studies yielded promising results, indicating that microbial U reduction could have potential as
an effective bioremediation tool. Following the same experimental scheme as the laboratory
sediment studies, field experiments were performed. This first in situ study was performed at
the Old Rife site in Colorado, where acetate was injected into the groundwater, and the levels
of Fe(III) and U(VI), as well as the microbial community were monitored (3). This field study (3),
as well as later experiments at the Oak Ridge FRC site in Tennessee (28, 47), reproduced results
of the lab-based experiments, showing the consumption of acetate concurrent with the
reduction of U(VI)/Fe(III), and the stimulation of the Geobacteracea.
More direct evidence specifically linking the metabolic activity of members of the
Geobacteraceae with U reduction was explored with the use of BioSep beads incorporating

13

C-

labeled acetate. Using stable isotope probing, the researchers were able to demonstrate that
the Geobacteraceae took up the labeled acetate, and dominated the microbial community in
the areas closes to the injection site, indicating that they are in fact metabolically active during
in situ bioremediation (8). Further evidence of metabolic activity was obtained by monitoring
the expression of a Geobacteraceae-specific citrate synthase gene (24). This gene is eukaryoticlike and phylogenetically distinct from that of other prokaryotes, but also a key component of
Geobacteraceae metabolism, making it an ideal candidate for monitoring the metabolic activity
5

of this family of organisms (24). When tested in lab and field experiments, the transcript levels
of the citrate synthase gene correlated well with the level of acetate, indicating that with the
addition of an electron donor, the Geobacteraceae were metabolically active (2, 24). Thus, they
are able to reduce U under conditions that are conducive to growth.

THE SEARCH FOR U(VI) REDUCTASES
Although U is not known to be an essential component for any enzyme or biological
structure (69), it has been shown that U can support the growth of Anaeromyxobacter
dehalogenans and Geobacter spp. (38, 62). When provided as an electron acceptor, however,
lower-than-expected growth yields were observed, potentially due to toxicity of the U (62). U is
known to be toxic to cells upon entering the cell envelope so it has been proposed that U
reduction takes place mainly outside of the cell (69). This is in agreement with observations that
precipitated U is often found to be localized outside of the cell, or in the periplasm (9, 21, 35,
41, 63, 69). It has therefore been suggested that the best candidates for U reductases are likely
to be located in the periplasm or outer membrane (35, 69).
As genome sequences became available, representatives of the Geobacteraceae were
found to have an unusually large number of genes coding for electron transfer proteins termed
cytochromes (45). Multi-heme c-type cytochromes are electron transfer proteins that are
responsible for passing electrons from the quinone/quinol pool to other cytochromes within
the cell membrane, or to exogenous electron acceptors (i.e. fumarate, Fe(III), U(VI)) (64). The
genome of the model organism for this family, Geobacter sulfurreducens, contains a total of
111 predicted c-type cytochromes (45). This unprecedented number may be an indication of
6

the versatility of electron acceptors that can be used by this organism, as well as the extreme
importance of the electron transfer process (45). The search for U reductases therefore focused
mainly on cytochromes present in the periplasm or outer membrane.
With the establishment of a genetic system in G. sulfurreducens (10) it became possible
to investigate the metabolic role and localization of individual cytochromes. In recent years
numerous studies in G. sulfurreducens have demonstrated the role of cytochromes in the
transfer of electron to electrodes, and the reduction of metals, including U(VI) (5, 26, 31, 33, 44,
54, 60, 63).
Studies to date have implicated periplasmic (36, 52, 63) as well as outer membrane (41,
63) cytochromes. U(VI) reduction was not completely eliminated by mutating the genes
encoding these cytochromes, however, and extensive periplasmic mineralization was observed
in all studies, suggesting that additional reductases remained active in the periplasm. These
putative periplasmic reductases are unlikely to be relevant to U reduction under environmental
conditions because, as discussed above, this situation is unlikely to be environmentally relevant
as it would be to the cell’s disadvantage to reduce U intracellularly.

PILUS NANOWIRES
It is important to note that these previous studies were carried out at temperatures that
prevent the expression of pilus nanowires (9), which are conductive extracellular appendages
required for the transfer of electrons to insoluble electron acceptors in the environment (57).
The G. sulfurreducens type IV pilus apparatus is very similar to those of other organisms in that
it utilizes the same set of core proteins including PilA (pilin subunit), PilD (prepilin peptidase),
7

PilC (inner membrane protein), PilB (assembly ATPase), PilT (retraction ATPase) and PilQ (outer
membrane secretin) (40, 42, 45). The PilA protein of G. sulfurreducens also possesses a
structure common to type IV pilins with a highly conserved N-terminal alpha-helical region, and
a more variable C-terminal region (42, 51, 65). Despite this conservation, the Geobacter pilin is
evolutionarily distinct from other pilins, and forms an independent line of descent (57). What
distinguishes the Geobacter pilin from that of other organisms is that it lacks the C-terminus
globular head domain commonly found in other pilins. For this reason, it has been predicted
that Geobacter pili may have adapted to perform a function distinct from that of other
organisms (57). This was confirmed by the observation that G. sulfurreducens pili were not
required for common functions such as adhesion to surfaces and twitching motility, but instead
were essential for the reduction of insoluble Fe(III) oxides (57). Conductivity measurements
performed with atomic force microscopy (AFM) demonstrated the high conductivity of the pilus
filaments, and implicated them as “pilus nanowires” which serve as the electrical conduit
between the cell and insoluble electron acceptors such as Fe(III) oxides (57).
As discussed above, Geobacter obtain most of their energy for growth during in situ U
bioremediation from Fe(III) oxides. As reduction of Fe(III) oxides requires the expression of the
pilus nanowires (57), we can infer that nanowires are also expressed during in situ
bioremediation of U. This hypothesis, coupled with the lack of a definitive extracellular U
reductase, led us to evaluate the contribution of the pilus nanowires to U(VI) reduction.
Chapter 2 of my dissertation describes a previously unrecognized role for Geobacter pili in the
extracellular reduction of U and their essential function as a catalytic and protective cellular
mechanism.
8

BIOFILMS IN THE SUBSURFACE
In addition to their role in electron transfer, the expression of pilus nanowires by G.
sulfurreducens also promotes cell-cell aggregation and biofilm formation (58). Biofilms are
defined as “matrix-enclosed bacterial populations adherent to each other and/or to surfaces or
interfaces” (12), and have long been observed in the environment, with the first reports dating
back to the 1930s and 40s when it was noted by several groups that “solid surfaces are
beneficial to bacteria” (72). It was not until many years later that this phenomenon was given a
name, and demonstrated to be both ubiquitous and the predominate mode of bacterial growth
in the environment (12-15, 20).
Upon encountering a surface for adhesion, bacteria undergo a variety of phenotypic
changes allowing them to attach to the surface, and develop a three-dimensional biofilm. These
phenotypic alterations can include the abundant production of exopolysaccharide (EPS), as well
as an increased resistance to heavy metals and other antibiotics (12, 23), with one study
demonstrating heavy metal resistance of the biofilm at levels up to 600-fold greater than that
of planktonic cells (66). Furthermore, it appeared that metal immobilization and toxicity was
localized to the exterior of the biofilm while the interior cells remained viable, potentially
because EPS is able to prevent the toxin from diffusing into the center of the biofilm (66).
This increased resistance of biofilms to toxic compounds led researchers to investigate
the potential for their use in bioremediation schemes. For example, biofilms of Pseudomonas
extremaustralis display higher growth and hydrocarbon degradation capabilities than
planktonic cells (68), and preliminary studies with yeast have shown promise in utilizing biofilms
9

as trickling filter for the degradation of diesel oil (7). Similarly, biofilm carriers were shown to be
an effective method of removing pharmaceutical substances (diclofenac, ketoprofen,
gemfibrozil, clofibric acid and mefenamic acid) from wastewater (16). In addition, it has been
hypothesized that U reduced by biofilms could be protected from reoxidation and
remobilization. While the U at the surface of the biofilm may become reoxidized as a result of
the oxygen intrusions common to the subsurface environment, it is possible that reducing
conditions will be maintained in the inner layers of the biofilm, thereby preventing reoxidation
of U (6). This is in contrast to cultures of planktonic cells from which bound U is readily oxidized
(6). Additionally, recent work has shown that following the addition of acetate to contaminated
groundwater Geobacter cells transition from growing predominately as planktonic cells to
dominating the metabolically active, sediment-associated biofilm community (30).
Despite these findings, the contribution of Geobacter biofilms to uranium removal and
reduction has not been investigated. Previous studies in planktonic cells have demonstrated the
role of the pilus nanowires, a cell envelope component essential to biofilm development, in U
immobilization and reduction (9). Thus, we sought to determine the role of pili in biofilm
development, as well as U reduction.
In chapter 3 of this dissertation I demonstrate that multilayer biofilms are able to reduce
and tolerate substantially more U than planktonic cells for prolonged periods of time (up to 24
h), making them an attractive option for the development of permeable biobarriers for U
bioremediation. I also demonstrate the role of pili as a U reductase in the biofilms.

10

BIOFILM DEVELOPMENT
The development of a biofilm progresses in several stages beginning with an approach
to the surface and initial attachment, colonization and microcolony development, and finally
growth into a mature biofilm (15, 48). The transition to each stage requires the expression of a
unique set of cellular components, many of them extracellular structures (22). For example,
motility proteins such as pili and flagella are often important for initial attachment and
colonization, while the EPS matrix facilitates the development of a mature biofilm (22, 70). This
is not representative of every organism, however, as some non-motile bacteria, such as
Staphylococcus epidermidis, utilize protein and/or polysaccharides as adhesins (22, 61).
The work presented in chapter 3 highlights the distinct role of each biofilm developmental
stage to the immobilization and reduction of U, leading us to examine the cellular components
characteristic of each stage. Previous studies have utilized random transposon mutagenesis
coupled with a high-throughput screening for biofilm-deficient phenotypes to identify genes
involved in the developmental process (39, 49, 50, 53, 56, 59, 67). Thus, to gain further insight
into how biofilms transform U I developed and screened a library of transposoninsertion mutants and identified mutants with biofilm defects. This study confirmed the role of
Geobacter pili in biofilm formation, and identified other cell envelope components involved in
biofilm formation and electron transport that may be relevant to U transformations in the
subsurface.

11

REGULATION OF CELL ENVELOPE COMPONENTS AND ELECTRON TRANSFER PROCESSES
The work described in the previous chapters of my dissertation has highlighted the
importance of the cell envelope and its components for the development of biofilms, as well as
the survival of Geobacter in the subsurface. However, little is known regarding the regulation
of the cell envelope. Thus, I investigated the role of the Geobacter’s extracytoplasmic function
(ECF) sigma factor, RpoE.
Sigma factors () such as RpoE are able to respond to environmental signals and direct the
bacterial RNA polymerase to specific promoters (71), allowing the cell to reprogram gene
expression in order to adapt to the given environment (27).
The ECF sigma factors, in particular, are responsible for coordinating the expression of
genes in response to environmental stressors, specifically those that cause imbalanced
synthesis of outer membrane components (43, 46). This can occur as a result of generalized
stressors such as temperature and ethanol (55), or more specific cues such as the transition
from exponential to stationary phase, or response to nutritional stress (11), as well as increased
sensitivity to adverse environmental conditions such as antibiotics, reactive oxygen species, and
changes in osmolarity (32).
In the last chapter of my dissertation, I show that RpoE is required for response to cell
envelope stress caused by oxygen, high pH, and low temperature, as well as the regulation of
Geobacter’s extracellular electron transfer pathways. This highlights the functional
specialization that RpoE has undergone to control the adaptive responses that enable
Geobacter bacteria to survive in the subsurface, and links my findings to Geobacter’s physiology
in the subsurface.
12

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13

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20

CHAPTER 2

EXTRACELLULAR REDUCTION OF URANIUM VIA GEOBACTER CONDUCTIVE PILI
AS A PROTECTIVE CELLULAR MECHANISM

The material presented in Chapter 2 has been adapted from the following publication:
Cologgi, D. L., S. Lampa-Pastirk, A. M. Speers, S. D. Kelly, and G. Reguera. 2011.
Extracellular reduction of uranium via Geobacter conductive pili as a protective cellular
mechanism. Proc Natl Acad Sci U S A 108:15248-15252.

21

ABSTRACT
The in situ stimulation of Fe(III) oxide reduction by Geobacter bacteria leads to the
concomitant precipitation of U(VI) from groundwater. Despite its promise for the
bioremediation of uranium contaminants, the biological mechanism behind this reaction has
remained elusive. Because Fe(III) oxide reduction requires the expression of Geobacter’s
conductive pili, we evaluated their contribution to uranium reduction in Geobacter
sulfurreducens grown under pili-inducing or non-inducing conditions. A pilin-deficient mutant
and a genetically complemented strain with reduced outer membrane c-cytochrome content
were used as controls. Pili expression significantly enhanced the rate and extent of uranium
immobilization per cell and prevented periplasmic mineralization. As a result, pili expression
also preserved the vital respiratory activities of the cell envelope and the cell’s viability.
Uranium preferentially precipitated along the pili and, to a lesser extent, on outer membrane
redox-active foci. In contrast, the pilus-defective strains had different degrees of periplasmic
mineralization matching well with their outer membrane c-cytochrome content. X-ray
absorption spectroscopy analyses demonstrated the extracellular reduction of U(VI) by the pili
to mononuclear U(IV) complexed by carbon containing ligands, consistent with a biological
reduction mechanism. In contrast, the U(IV) in the pilin-deficient mutant cells also required an
additional phosphorous ligand, in agreement with the predominantly periplasmic
mineralization of uranium observed in this strain. These results demonstrate a previously
unrecognized role for Geobacter conductive pili in the extracellular reduction of uranium and
highlight its essential function as a catalytic and protective cellular mechanism that is of interest
for the bioremediation of uranium-contaminated groundwater.
22

INTRODUCTION
Dissimilatory metal-reducing microorganisms gain energy for growth by coupling the
oxidation of organic acids or H2 to the reduction of metals. Some can also use U as an electron
acceptor (42, 58), a process that could be harnessed for the bioremediation of the
contaminated aquifers and sediments left by the intensive U mining practices of the Cold War
era (39). Interestingly, stimulating the activity of metal-reducing microorganisms in situ resulted
in the concomitant removal of soluble U (U(VI)) from the contaminated groundwater and
detection of its sparingly soluble, less mobile form, U(IV) in sediments (1, 9, 25, 51, 73). This
suggests that stimulating metal reduction in the subsurface results in the biological reduction of
U(VI) to U(IV), thereby preventing plume migration and eliminating the potential for
contaminant exposure.
The removal of U(VI) from groundwater following the in situ stimulation of metal reduction
is often concomitant with substantial increases in the growth and activity of dissimilarity metalreducing microorganisms in the family Geobacteraceae (1, 9, 51, 72). Despite extensive efforts
to understand the mechanisms and pathways used by these bacteria to reduce U(VI), the
nature of its U reductase has remained elusive. Early studies with Geobacter metallireducens
GS15 suggested that U was reduced extracellularly to uraninite under conditions conducive to
cell growth (17, 42). The development of genetic tools in Geobacter sulfurreducens (12)
motivated molecular studies to elucidate the biological mechanism behind this reaction.
Because c-cytochromes are abundant in the cell envelope of Geobacter bacteria, studies
focused on identifying extracytoplasmic c-cytochromes that could function as dedicated U
reductases (38, 63). However, mutations were often pleiotropic (31-33) and either showed no
23

defect or only partial defects in the cell’s ability to remove U(VI) (38, 63). Interpretation was
also difficult due to inconsistencies in the reported mutant phenotypes, with some mutations
reportedly abolishing U(VI) removal activities yet mutant cells showing extensive mineralization
(63). Furthermore, these studies consistently showed that the U precipitated inside the cell
envelope. U is not known to be essential for the synthesis of any cell component or for any
cellular biological reaction yet can be reduced and precipitated nonspecifically by the abundant
low potential electron donors of the cell envelope of Gram-negative bacteria (71). This is
predicted to compromise the integrity of the cell envelope and its vital functions (64). Because
of this, the environmental relevance of these early studies in G. sulfurreducens is questionable.
The energy to support the growth of Geobacter bacteria after in situ stimulation results
from the reduction of the abundant Fe(III) oxides, a process that requires the expression of
their conductive pili (55). In contrast to the lack of conservation of c-cytochrome sequences (8),
the genes encoding the Geobacteraceae pilus subunits or pilins are highly conserved and form
an independent line of descent (55). This is consistent with the pili’s specialized function as
electrical conduits. The pilus apparatus is anchored in the cell envelope of Gram-negative cells
(49, 67, 70) and could potentially accept electrons from cell envelope c-cytochromes or the
menaquinone pool in the inner membrane. Pili also protrude outside the cell envelope and can
reach m lengths, thereby enhancing the cell’s reactive surface. Thus, we hypothesized that the
pili could catalyze the reduction of U(VI) ‘at a distance’ to maximize the cell’s catalytic surface
while minimizing exposure to U. Here we show that the conductive pili of G. sulfurreducens
catalyze the extracellular reduction of U(VI) to a mononuclear U(IV) phase and prevent its
periplasmic mineralization, thereby preserving the functioning and integrity of the cell envelope
24

and the cell’s viability. These results identify pili as the primary U reductase of Geobacter
bacteria and demonstrate that their catalytic function also serves as a cellular protective
mechanism. This suggests that their expression confers on Geobacter bacteria an adaptive
ecological advantage in the contaminated subsurface of potential interest for the optimization
of in situ bioremediation.

MATERIALS AND METHODS
Strains and culture conditions. Wild-type (WT) Geobacter sulfurreducens PCA (ATCC
-

51573), a pilin-deficient mutant (PilA ) (55), and its genetically complemented strain
(pRG5::pilA) (55) were routinely revived from frozen stocks in NB medium (12) supplemented
with 15mM acetate and 40mM fumarate (NBAF) and incubated at 30 °C. The cultures were
transferred to fresh water (FW) medium (41), prepared with the modifications described below,
and supplemented with 15mM acetate and 40mM fumarate (FWAF). The electron donor and
acceptor were prepared as autoclaved concentrated stocks (0.75 M sodium acetate and 1 M
sodium fumarate at pH 7, respectively). FW medium was prepared from a concentrated (10x)
basal FW medium stock containing NaHCO3 (25 g/L), NaH2PO4·H2O (0.6g/L), NH4Cl (2.5 g/L),
and KCl (1.0 g/L). Vitamins were prepared as separate solutions as previously described (3).
Trace minerals were prepared as previously described (40), except that ZnSO4 was replaced
with ZnCl2 (0.13 g/L), and Na2WO4·2H2O (0.025 g/L) was added. FWAF medium contained FW
stock (96 ml/L), 0.75 M sodium acetate (20 ml/L), 1 M sodium fumarate (40 ml/L), vitamin
25

solution (10 ml/L), mineral solution (10 ml/L). The medium was dispensed in pressure tubes or
serum bottles, sparged with N2:CO2 (80:20) to remove dissolved oxygen and sealed with butyl
rubber stoppers and aluminum tear off seals (Wheaton) prior to autoclaving. Unless otherwise
indicated, all cultures were incubated at 30°C.
Pili induction at suboptimal growth temperatures. WT controls expressing pili (WTP+)
were obtained by growing WT cells at suboptimal growth temperatures (25 °C). Bacterial pili
expression is often thermoregulated (16), an adaptive mechanism that enables bacteria to
rapidly assemble the pili in environments where pili functions are advantageous (65). As in
other bacteria (57), pili production in Geobacter is induced at suboptimal growth temperatures
(25°C) mimicking the suboptimal growth conditions that the cells encountered during the
reduction of Fe(III) oxides (55). In contrast, growth at 30°C prevented pili assembly and
produced a pili-deficient strain WTP-.
U(VI) resting cell suspension assays. Resting cell suspensions were prepared as
described elsewhere (63), except that cells were harvested from mid-log phase cultures (OD600,
0.3-0.5) and resuspended in 100 ml reaction buffer with 20 mM sodium acetate to a final OD 600
of 0.1. Heat-killed controls were prepared by autoclaving the cultures for 30 min. Suspensions
were incubated for 6 h at 30°C with 1 mM uranyl acetate (Electron Microscopy Sciences), as
previously described (63). After incubation, 500 l samples were withdrawn, filtered (0.22-µm
Millex-GS filter, Millipore) to separate the cells, acidified in 2% nitric acid (500 l), and stored at
26

-20°C. All procedures were performed anaerobically inside a vinyl glove bag (Coy Labs)
containing a H2:CO2:N2 (7:10:83) atmosphere. The initial and final concentration of U(VI) in the
acidified samples was measured using a Platform Inductively Coupled Plasma Mass
Spectrometer (ICP-MS) (Micromass, Thermo Scientific) to calculate the amount of U(VI)
removed from solution.
Gene expression analyses by qRT-PCR. Quantitative reverse-transcriptase PCR (qRTPCR) was used to quantify pilA and recA transcripts in RNA extracted from resting cells of the
WTP+ and WTP- strains before and after incubation with uranyl acetate for 6 h. WT P+ controls
incubated in the same reaction buffer without uranyl acetate also were included. RNA was
extracted using the TRIzol reagent (Invitrogen, Carlsbad, CA), and treated with RQ1 RNase-free
DNase (Promega, Madison, WI) according to the manufacturer’s instructions. Reverse
transcription was performed using Superscript III Reverse Transcriptase (Invitrogen, Carlsbad,
CA) following manufacturer’s recommendations. Primer pairs RT_ORF02545_F and
RT_ORF02545_R were used for pilA (13) and recA660f and recA737r were used for recA (20).
For qRT-PCR, the cDNA generated after reverse transcription was diluted 1:1000 in a 25 µl
reaction that contained each primer (5 µM) and SYBR Green MasterMix (Applied Biosystems,
Foster City, CA), according to the manufacturer’s instructions. Samples were amplified using a
Bio-Rad (Hercules, CA) iCycler (iQ5 Multicolor Real-Time PCR Detection System). The
comparative CT method (61) was used to calculate the relative expression of the pilA gene using
the recA constitutive expression as internal control. Briefly, the ∆CT value (CT (pilA) – CT (recA))

27

was calculated for triplicate biological replicates before (0 h) and after (6 h) incubation with
U(VI) acetate. The average of the difference between the 6 h and 0 h CT values was used to
calculate the ∆∆CT. The relative fold change in pilA expression versus the recA internal control
was calculated with the formula 2

-∆∆CT

.

TEM and Energy Dispersive Spectroscopy (EDS) analyses. After U exposure, resting
cells were adsorbed onto 300-mesh carbon-coated copper grids (Ted Pella), fixed with 1%
glutaraldehyde for 5 min, and washed 3 times with ddH2O for 2 min. Unstained cells were
directly imaged with a JEOL100CX electron microscope (Japan Electron Optic Laboratory)
operated at a 100 kV accelerating voltage. Imaging and elemental analysis of the extracellular
uranium precipitates was performed with a JEOL 2200FS operated at 200 kV and an Energy
Dispersive Spectroscopy (EDS) detector.
For thin sections, cells were harvested by centrifugation (1,200 x g, 30 min, room
temperature) and prepared as previously described (43), except that a Power Tome XL (RMC
Products, Boeckler Instruments, Tucson, AZ) was used for sample sectioning. Thin sections were
imaged with a JEOL100CX operated at 80 kV. Approximately 400-500 cells from randomly
taken, thin-sectioned fields were also examined for periplasmic mineralization. A cell was
considered to have a mineralized periplasm when it contained both a fully mineralized outer
membrane and generalized mineralization in the periplasm and/or inner membrane.
Pili purification, quantification, and biochemical characterization. Pili were purified as
detergent-insoluble fractions from cells first lysed by sonication and treated with RNase, DNase

28

and lysozyme enzymes (11). Three biological replicates were used for each strain. Cell
membranes and proteins in the extracts were solubilized with sodium dodecyl sulfate (SDS, 1%
final concentration) and separated by preparative 12% polyacrylamide gel electrophoresis (5%
stacking gel, 40 mA for 5 h) using a Prep Cell 491 apparatus (Bio-Rad). The detergent-insoluble
fraction, which did not enter the stacking gel, was recovered by aspiration, washed in ddH 2O,
extracted once with 95% ethanol (Decon Laboratories), and dried in a SpeedVac system (Savant
Instruments Inc.) at room temperature for 20 min. The dry sample was resuspended in 1 ml of
ddH2O and vortexed for 60 seconds to break up large clumps before extracting poorly bound,
soluble protein with 0.2 M glycine (pH 1.5, adjusted with HCl; Invitrogen) at 100°C for 10 min.
The insoluble fraction was recovered by centrifugation (16, 000 x g, 25 min, 4°C), washed five
times with ddH2O, and dried in a SpeedVac at room temperature prior to storage at -20°C. The
amount of pili protein was measured after resuspending the dry samples in 10 mM CHES buffer
(pH 9.5), incubating at 4°C overnight, and mixing 1:1 with the working reagent solution of the
Pierce Microplate BCA protein assay kit (reducing agent compatible, Thermo). The samples
were incubated at 60°C for 1h before spectrophotometric measurements. BSA was used as a
standard.
For PAGE analyses, dried preparations of purified pili were resuspended in 5 μl of ddH2O
containing 10% (w/v) Octyl-β-D-Glucopyranoside (OG) (Sigma, 98%) and incubated at room
temperature for 2 h. The samples were diluted with 5 volumes of ddH2O to reduce the OG

29

concentration to 2% (v/v) and incubated for an additional 24 h period at room temperature.
The OG-treated sample was boiled in SDS-PAGE sample buffer (59) and subjected to
electrophoresis through a 10-20% Tris/Tricine ReadyGel (Bio-rad) using a Mini Protean Tetra
Cell apparatus (Bio-Rad). After electrophoresis, the gels were silver stained using the Pierce®
Silver Stain for Mass Spectrometry kit (Thermo Scientific), following the instructions supplied by
manufacturer, and scanned. The migration of the molecular mass standards in the gradient gel
2

was estimated and fitted a polynomial distribution (R = 0.95523), which was used to calculate
the molecular mass of unknown bands. Duplicate gels were also electrophoretically transferred
to a PVDF membrane (HyBond LFP, Amersham GE Healthcare) at 25 V for 150 min using a Mini
Protean Tetra Cell apparatus (Bio-rad). The Amersham ECL Plex Western blotting kit was used
for the electrophoretic transfer and membrane blocking, following manufacturer’s
recommendations. After blocking, the membrane was incubated in 10 mL antibody diluent
solution (90 min, room temperature, gentle agitation) containing a 1:5,000 dilution of the
primary antibody (rabbit α-PilA polyclonal antibodies raised against the 42 C-t amino acids of G.
sulfurreducens PilA protein) and a 1:2,500 dilution of goat α-rabbit IgG antibodies conjugated to
the Cytm 5 fluorescence dye (ECL™ Plex, Amersham GE Healthcare). The membrane was
washed 4 x 5 minutes in wash buffer (TBS-T, pH 7.4, 0.1% Tween 20) and rinsed three times in
wash buffer without Tween 20. The protein bands that hybridized with the primary antibodies
were visualized after scanning the membrane blot with a Typhoon imager operated in
fluorescence mode (excitation at 633 nm, 670 BP 30 filter, and PMT setting at 600 V). SDS-PAGE
and western blotting experiments were performed by Dr. Sanela Lampa-Pastirk.
30

Transmission Electron Microscopy (TEM) and Confocal Laser Scanning Microscopy
(CLSM) analyses of purified pili. Detergent-insoluble fractions were examined by TEM and
CLSM. For TEM, an aqueous solution of purified pili was adsorbed on a carbon-copper grid (300
Mesh, Electron Microscopy Sciences), negatively stained with 1% (w/v) uranyl acetate in
distilled water, and examined with a JEOL 100 CX electron microscope operated at 100 kV. For
CLSM, detergent-insoluble fractions were dissolved in phosphate buffer saline (PBS, pH 7),
adsorbed onto glass coverslips for 30 min, washed with PBS and fixed with 3.7%
paraformaldehyde in PBS. The samples were then incubated for 30 min in PBS-1% BSA, before
overnight incubation at 4°C with a 1:100 dilution anti-PilA primary antibody in PBS-1% BSA.
After three washes in PBS-1% BSA, the samples were incubated with a 1:1,000 dilution of antirabbit antibodies conjugated to the Alexa fluor 488 dye (Molecular Probes, Invitrogen) for 1 h.
The coverslips were then washed 3 times with PBS and examined with a Zeiss LSM Pascal
confocal microscope equipped with a Plan-Neofluar 63x oil objective (excitation, 488 nm;
emission, 505-535 nm). Visualization and antibody detection of purified nanowires was
performed by Dr. Sanela Lampa-Pastirk.
SDS-PAGE and staining of outer membrane, heme-containing proteins. Outer
membrane c-cytochromes were mechanically detached from all the strains and isolated as
previously described (47). A mutant deficient in the outer membrane c-cytochrome OmcS (47)
and grown at 25°C was included as a control. Proteins (2.5 μg) in the supernatant fluids were
separated by Tris-Glycine denaturing polyacrylamide gel electrophoresis (2) using a 12% MiniProtean TGX gel (Bio-Rad). The Novex Sharp molecular weight markers (Invitrogen) were used

31

as standards. Reducing agents were omitted from the SDS-sample buffer and the samples were
loaded onto the gel without boiling to prevent the loss of heme groups (68). C-type
cytochromes were detected as heme-stained bands using N,N,N′,N′-tetramethylbenzidine, as
previously described (68). After heme-staining, the gels were photographed, destained with 70
mM sodium sulphite (68), and silver-stained for total protein with the Pierce silver stain for
Mass Spectrometry (Thermo).
X-ray Adsorption Spectroscopy (XAS) analyses. Resting cells incubated with U for 6 h
were harvested by centrifugation (13,000xg, 10 min, 4°C) and loaded into custom-made plastic
holders, triply packaged in Kapton film and sealed with Kapton tape (DuPont), under anaerobic
conditions. Samples were stored at -80°C and kept frozen during XAS measurements, which
were performed with a multielement Ge detector in fluorescence mode, using the PNC-CAT
beamline 20-BM at the Advanced Photon Source (Argonne National Laboratory) and standard
beamline parameters, as described elsewhere (29). XANES measurements were used to
calculate the relative amount of U(VI) to U(IV) by linear combination fitting of the spectrum
with U(VI) and U(IV) standards. The spectra were energy aligned by simultaneously measured
uranyl nitrate standard. The uranium EXAFS spectra were processed using the methods (48) in
Athena (54) and were modeled using FEFFIT (66) with theoretically generated models from FEFF
7.02 (74), as described elsewhere (29). Multiple scattering paths from distant C3 and Odist
atoms were also included in the model, yet did not have a significant contribution. The
coordination number for U-Oax (Noax values) obtained from EXAFS measurements of 3 to 4
biological replicates from at least two independent experiments also was used to estimate the
32

amount of U(VI) reduced to U(IV) in each sample. For example, a Noax value of 0.3 was
obtained for one of the biological replicates of the WTP+ (as given in Table 2.2). 100% of U(VI)
would have a U-Oax coordination number of 2.0. Therefore the amount of U(IV) that
corresponds to the WTP+ Noax is 0.3/2.0 = 0.15. This indicates that there is 15% U(VI) and 85%
U(IV) in this sample. Analysis of all data generated through XANES and EXAFS analyses was
performed by Dr. Shelly Kelly.
Vitality and viability assays. The RedoxSensor Green reagent (Molecular Probes) was
used to measure the cell’s vitality (broadly defined as the levels of activity of cell’s vital
reactions) after U exposure. This reagent yields green fluorescence when modified by the
bacterial reductases, which are mostly located in the electron transport system of the cell
envelope (19, 26). Resting cells were harvested in a microcentrifuge (12,000xg), washed, and
resuspended in a 100 l Tris-Buffered Saline (TBS) before mixing it with an equal volume of a
working concentration of the dye. Fluorescence was measured in two biological replicates, with
two technical replicates each, using a SpectraMax M5 plate reader (Molecular Devices) with an
excitation of 490nm and emission of 520nm. Cell viability after U exposure in comparison to
controls without U was assayed by recovering the resting cells in NBAF medium and measuring
the length of the lag phase, as described before (36). Prior to inoculation, resting cell
suspensions were gassed for 15 min with filtered-sterilized air to reoxidize the U deposits (63)
without compromising G. sulfurreducens viability (36). Cells were harvested by centrifugation
(1,200 x g, 5 min), resuspended in 1 ml of wash buffer (final OD600 of 0.4), and mixed with 10

33

ml of NBAF in pressure tubes. The cultures were incubated at 25°C and growth was periodically
monitored as OD600.

RESULTS
Expression of pili promotes the extracellular reduction of U(VI). The correspondence
between pili expression and U immobilization was examined by monitoring the removal of
U(VI) acetate from solution by resting wild-type cells incubated at 25°C (WTP+) or 30°C (WTP-)
-

to induce or prevent pili assembly, respectively. Controls with a pilin-deficient mutant (PilA )
and its genetically complemented strain (pRG5::pilA) were also included. The piliated strains
WTP+ and pRG5::pilA removed substantially more U(VI) from solution than the nonpiliated
-

strains WTP- and PilA (Fig. 2.1, top).

34

FIGURE 2.1: Reduction and subcellular localization of U. Reduction of U(VI) to U(IV) (top) and
TEM images (bottom) of unstained whole cells showing the subcellular localization of the U
-

deposits in the WTP+, WTP-, PilA , and pRG5::pilA strains (left to right). Bar, 0.5 µm.

The activity was biological in nature, as heat-killed WTP+ and WTP- controls did not remove
U(VI) significantly (0.02  0.04 and 0.05  0.02 mM, respectively). X-ray Absorption Near Edge
Structure (XANES) spectroscopy confirmed the reductive nature of the U removal activity and
measured an average of 70-85% U(IV) in all samples (Fig. 2.1, top). Furthermore, the expression
of the pilA gene relative to the internal control recA did not change during the assay (Fig. 2.2),
thus ruling out any de novo pilin transcription.
35

Relative fold change (2-ΔΔCT)

5
4
3
2
1
0
-1

WTP+

WTP-

WTP+ (no U)

FIGURE 2.2: Fold change expression of pilA relative to recA in resting cell suspensions of the
WTP+ and WTP- strains after 6 h of incubation with 1 mM U(VI) acetate. A WTP+ control
incubated in reaction buffer without U(VI) acetate also is shown (WT P+ (no U)).

The extent of U(VI) removal also corresponded well with the levels of piliation, which were
measured as the protein content of purified PilA-containing pili samples (Fig. 2.3).

36

B

A

FIGURE 2.3: Micrographs of purified pilus fibers. TEM (A) and CLSM (B) micrographs of,
respectively, negatively-stained and anti-PilA immunodetected pilus fibers (displayed as green)
isolated as detergent-insoluble fractions in the WTP+. Inset in (A): SDS-PAGE Tricine gel (left)
and Western blot using anti-PilA polyclonal antibodies (right) showing the depolymerization of
the purified pili into the PilA peptide subunits. Numbers at left are molecular mass standards in
KDa and were used to estimate the apparent mass of the PilA band (6.6 kDa, as predicted for
the mature PilA based on amino acid sequence). For interpretation of the references to color in
this and all other figures, the reader is referred to the electronic version of this dissertation.

The level of pRG5::pilA piliation (3.6  1.7 g pili/OD600) was 2.5-fold higher than WTP+ (1.5 
0.1 g/OD600), which matched well with its superior capacity to remove U(VI) from solution
-

(1.8  1.0-fold higher than WTP+). By contrast, WTP- and PilA samples had no detectable pili
protein and reduced less U(VI).
The location of the U reductase system was studied by examining the cellular
localization of the U deposits in unstained whole cells by TEM (Fig. 2.1, bottom). The piliated
strains, WTP+ and pRG5::pilA, preferentially deposited the U extracellularly and in a
monolateral fashion, consistent with the localization of Geobacter’s conductive pili to one side

37

of the cell (55). The pili filaments were interspersed with the dense deposits (Fig. 2.4, inset).
Elemental composition analyses of the pili-associated deposits by TEM-EDS in the WTP+
confirmed the presence of U (Fig. 2.4).

10 nm

FIGURE 2.4: Energy dispersive X-ray spectrum of the pili-associated electron-dense deposits
imaged by TEM. Uranium peaks are highlighted with red arrows. Inset shows pili filaments (red
arrows) interspersed with the electron-dense uranium deposits. The text in this figure is not
meant to be readable, but is for visual reference only.
-

In contrast, extracellular U mineralization in the non-piliated strains, WTP- and PilA was limited
to the cell’s outer membrane and to membrane vesicles. TEM thin sections of the unstained
cells confirmed the presence of extracellular, needle-like U deposits in the piliated strains as
well as discrete regions of U deposition on the outer membrane (Fig. 2.5).

38

WTP+

WTP-

PilA-

pRG5::pilA

FIGURE 2.5: Subcellular localization of uranium deposits by TEM analyses of thin sections of
-

pili-expressing (WTP+ and pRG5::pilA) and non-expressing (WTP- and PilA ) cells. Top panels
show large fields (bar, 500 nm) and bottom panels show close-ups of a representative cell (bar,
100 nm). Needle-like extracellular precipitates are noticeable in the top micrographs of the
piliated strains, which is consistent with pili-associated uranium deposits. Discrete uranium
deposits on the outer membrane are also noticeable in most cells of the WT P+ and WTP- and in
a few cells of the pRG5::pilA strain, while fully mineralized outer membranes and periplasmic
-

deposition is the observed in most PilA cells (top and bottom panels). Mineralized membrane
-

vesicles budding from the cell (WTP-) or detached (PilA ) are indicated with red arrows.

Only a few cells (8  3% of the WTP+ and <1% of the pRG5::pilA) had periplasmic mineralization.
Outer membrane foci of U deposition were also noticeable in the WTP- cells, but more cells (37
 13% of the cells) had periplasmic deposition. The increased periplasmic mineralization in the
WTP- cannot be attributed to a differential expression of outer membrane c-cytochromes, as

39

outer membrane protein fractions had the same heme profile and content as the WT P+ (Fig.
2.6).

A

B
260
110
60
50
40
30
20
10

1

2

3

4

5

1

2

3

4

5

FIGURE 2.6: SDS-PAGE profiling of mechanically detached outer membrane proteins. Samples
were resolved in 12% (wt/vol) SDS-PAGE gels and silver-stained for total protein (A) and for
heme content (B). Approximately 2.5 µg of protein were loaded per lane. Lanes: 1, WT P+; 2,
-

-

WTP-; 3, PilA mutant; 4, pRG5::pilA; and 5, OmcS mutant. Numbers at right are molecular
weight standards in kilodaltons. The migration of the OmcS c-cytochrome is marked with an
arrow.
-

Periplasmic mineralization was more pronounced (85  12% of the cells) in the PilA mutant
(Fig. 2.5), which was also partially defective in outer membrane c-cytochrome production (Fig.
2.6). This suggests that outer membrane c-cytochromes also contribute to the extracellular
reduction of U(VI). It is unlikely that the outer membrane c-cytochrome OmcS, which has been
hypothesized to mediate electron transfer between the conductive pili and metals (35),
contributed to the pili-mediated reduction of U, because the pRG5::pilA strain expressed OmcS

40

at wild-type levels (Fig. 2.6) yet reduced more U than the WTP+ (Fig. 2.1A) and proportionally to
the levels of conductive pili assembled. Furthermore, the pRG5::pilA strain also had a defect in
outer membrane, heme-containing proteins (Fig. 2.6), yet cells had very little U deposition in
their cell envelope (Fig. 2.5). This is consistent with the pili functioning as the primary site for U
reduction.
X-ray absorption fine structure (EXAFS) analyses demonstrate the reduction of U(VI) to
mononuclear U(IV). U LIII-edge EXAFS spectra were modeled to determine the atomic
coordination about U and characterize the U(IV) product in all the strains (28). Models for the
EXAFS spectra included signals from neighboring P, U, and Fe atoms, but only C neighbors were
found to accurately reproduce the measured spectra. The spectra were best described by a
-

mixture of U(IV) and U(VI) coordinated by C-containing ligands. Only the PilA mutant required
an additional P ligand. A U signal corresponding to the U-U distance in uraninite at 3.87 Å was
tested but was inconsistent with the measured spectra. Fig. 2.7 shows the magnitude of the
Fourier transformed spectra and models for each spectrum.

41

FIGURE 2.7: U LIII-edge EXAFS spectra and models. (A) Magnitude of Fourier transform spectra are offset for clarity. Real part of
Fourier transform of WTP+ (B) and PilA (C). The components of the model are shown offset beneath the total model. Insets in B and
C show the U(IV) moiety that is consistent with the measured EXAFS spectra (U(IV), light grey; O, red; C, black; P, dark grey)

42

Fig. 2.7B and 2.7C show, as examples, the contribution of each path in the model in the real
-

part of the Fourier transform for the WTP+ and PilA cells and insets show a molecular moiety of
the U(IV) atomic environment that is consistent with the measured EXAFS. The WT P+ model
includes one C ligand bound to two O atoms of U in a bidentate fashion and followed by a
distant C atom (C3) and another C ligand bonded to one O atom of U(IV) in a monodentate
fashion and attached to a distant O atom (Odist). This model was simultaneously refined to all
-

spectra and was insufficient to reproduce the PilA spectrum, which required an additional
2

monodentate P ligand (Fig. 2.7C). The distances and  values used to model the spectra are
listed in Table 2.1. The coordination numbers (Table 2.2) are consistent with 1 to 2 bidentate C
ligands and 2 monodentate C ligands per U atom.

43

2

TABLE 2.1: EXAFS modeling results for R and σ

*
2

3 2

Path

CN

R (Å)

σ (∙10- Å )

Oax
Oeq
C1
C2

1.79 ± 0.01
2.37 ± 0.01
2.94 ± 0.01
3.43 ± 0.01
3.58 ± 0.01

2
15 ± 1
5±2
5±2

Oax1-Oax2

Noax
Noeq
Nc1
Nc2
Noax

Oax1-U-Oax2

Noax

3.58 ± 0.01

4

Oax1-U-Oax1

2Noax

3.58 ± 0.01

8

P1*
C3
C1-C3
C1-C3-C1
Odist
C2-Odist
C2-Odist-C2
CN, coordination number.

Np1
Nc1
2Nc1
Nc1
Nc2
2Nc2
Nc2

3.57 ± 0.05
4.41 ± 0.02
4.41 ± 0.02
4.41 ± 0.02
4.55 ± 0.02
4.58 ± 0.02
4.60 ± 0.02

5±2
5±2
5±2
5±2
5±2
5±2
5±2

†

†

4

†
†
†

Value held.

*PilA- mutant data set only.
TABLE 2.2: EXAFS modeling results for coordination numbers
Data set
Noax
Noeq
C1
P1
C2
0.3 ± 0.1 7.6 ± 0.4 1.8 ± 0.3
2.5 ± 0.4
WTP+
WTP-

0.4 ± 0.1

7.5 ± 0.3 1.7 ± 0.3

-

2.5 ± 0.4

PilA
pRG5::pilA

0.9 ± 0.1
0.8 ± 0.1

7.1 ± 0.4 2.0 ± 0.4
7.1 ± 0.4 1.6 ± 0.3

0.5 ± 0.3

2.3 ± 0.5
2.3 ± 0.5

-

The number of Oax atoms (Noax) was also used to estimate the amount of U(IV) in these
samples, as there are two Oax atoms for each U(VI) atom and none for U(IV) (28). An average of
3-4 replicates for each strain gives the values of 72 ± 16% (WTP+), 81 ± 6 % (pRG5::pilA), 85 ± 5%
-

(WTP-), and 76 ± 10% (PilA ). This provides additional evidence that, while the extent of U(VI)
44

removal depended on the expression of the pili, the ability of the cells to reduce the
immobilized U(VI) to U(IV) did not.
U reduction via pili as a cellular protective mechanism. The reverse correlation
between piliation and periplasmic mineralization suggested that the pili-mediated reduction
prevented U from permeating and being reduced in the periplasm, thus preserving vital
functions of the cell envelope. To test this, we used the fluorogenic RedoxSensorTM Green dye
to measure the vitality (or levels of vital reactions of the cell, in this case the cell’s reductase
activity) of the strains after U exposure in reference to unexposed controls. The vitality
remaining after U exposure was higher in the piliated strains and proportional to the levels of
piliation (pRG5::pilA>WTP+) (Fig. 2.8A).

45

FIGURE 2.8: Effect of U(VI) exposure on cell vitality and viability. (A) Vitality was measured as
bacterial reductase (respiratory) activity with the RedoxSensor dye in resting cells of the pili-

expressing (WTP+ and pRG5::pilA) and non-expressing (WTP- and PilA ) strains and expressed as
+

-

the ratio of relative fluorescence units emitted by from cells incubated with (U ) or without (U )
U. (B) Growth recovery of resting cells of the pRG5::pilA (black), WTP+ (dark gray), and WTP(light gray) after 6 h of U exposure (circles) in comparison to controls without U (lines).

Inasmuch as the RedoxSensor dye can also serve as a proxy for the integrity of the cell envelope
and the cell’s viability (19), these results suggested that the cell viability was also higher in the
piliated strains. To test this, we recovered the resting cells in growth medium and studied the
cell’s survival (defined as the cell’s ability to maintain its integrity and undertake division (4))

46

after exposure to U as a function of the length of the lag phase (Fig. 2.8B). While cells that had
not been exposed to U recovered rapidly and simultaneously, the strains exposed to U
recovered in a step-wise fashion. The lag phase was shortest (~ 18 h) in the hyperpiliated
pRG5::pilA cells, followed by the WTP+ (~56 h) and the WTP- (~81 h), and correlated well with
2

-

the levels of periplasmic mineralization of the strains (R = 0.947). The PilA mutant recovery
was similar to the other non-piliated strain, WTP-, yet more variable (lag phases ranging from
72 to 82 h). It also grew faster (~9 h doubling time compared to ~11 h for the WT and
pRG5::pilA strains) than the other strains, which is expected to accelerate recovery. Despite
these differences, the survival rates (calculated as the reverse of the length of the lag phase) of
2

all the strains followed a linear regression (R = 0.908) with the levels of pili protein.

DISCUSSION
Physiological relevance of the extracellular reduction of U by Geobacter’s pili. Our
results show that cells that assembled pili immobilized a greater amount of U and also
prevented it from permeating inside the periplasm, where it would have otherwise been
reduced nonspecifically by c-cytochromes and other low potential electron donors (71). As a
result, the extracellular reduction of U via pili also preserved the vital functions of the cell
envelope and the cell’s viability. This mechanism is consistent with field studies showing that
the indigenous Geobacter community that is stimulated during in situ bioremediation is
metabolically active (23, 72) and gains energy for growth from the reduction of Fe(III) oxides (9,

47

14), a process that requires the expression of the conductive pili (55). We used a temperaturedependent regulatory switch (see Materials and Methods) to produce WT controls (WTP-) that
did not assemble pili, yet had WT levels and profiles of outer membrane cytochromes. The lack
of pili in the WTP- strain significantly diminished the cell’s ability to reduce U(VI), increased the
degree of periplasmic mineralization, and reduced the respiratory activity of the cell envelope
and the cell’s viability. WTP- cells also had extensive outer membrane vesiculation, a process
linked to the selective detoxification of unwanted periplasmic materials by cells undergoing cell
-

envelope stress (45). Similarly, the inability of a PilA mutant to produce pili impaired the yields
of U reduction. This mutant strain also had reduced outer membrane cytochrome content and,
as a result, more U traversed the outer membrane and precipitated in the periplasm. The fact
that the pili of G. sulfurreducens catalyze the extracellular reductive precipitation of U under
physiological conditions conducive to growth, is also in agreement with early studies with G.
metallireducens suggesting that the reduction of U is extracellular (17) and coupled to cell
growth (42). In these studies (17, 42), cells were grown with Fe(III) citrate as an electron
acceptor, which are culture conditions that promote pili expression in G. metallireducens (10)
but not in G. sulfurreducens (55). Thus, the extracellular precipitation and sustained removal of
U reported for G. metallireducens is consistent with pili catalyzing the reaction as well.
Reduction of U to mononuclear U(IV) phases. Despite differences in the mechanism
and yields of U reduction, the strains with the lowest levels of periplasmic mineralization
(WTP+, pRG5::pilA and WTP-) produced similar U LIII-edge EXAFS spectra that were modeled as

48

mostly U(IV) coordinated by C-containing ligands in bidentate and monodentate fashion and
that lacked any Fe- or P-containing ligands. The bidentate C1-C3 ligand is likely biological in
nature as reported for the carboxyl coordinations involving amino acids and lipolysaccharide
-

sugars (5, 6, 37). In contrast, the PilA mutant, which had the highest degree of periplasmic
mineralization, required an additional monodentate P ligand. This signal was small with a
coordination number of 0.5 ± 0.3, indicating that, on average, 50% of the U atoms contained a P
ligand while the other 50% shared the atomic coordination of the other strains. Alternative
interpretations such as 25% or 12.5% of U atoms with 2 or 4 P ligands, respectively, are unlikely
because the CN values for the C1 and C2 signals did not decrease proportionally (50% and 75%,
-

respectively). It is also unlikely that the low levels of U(VI) reduced by the PilA strain
contributed to the distinct spectra, because WTP- cells reduced less U(VI) and did not require a
P ligand for U coordination. The P coordination and the generalized periplasmic mineralization
-

observed in the PilA mutant cells suggest that U(VI) permeated deep and fast into the cell
envelope, where it formed carboxyl and phosphoryl coordinated complexes with periplasmic
proteins and the peptidoglycan layer (5, 6) and membrane phospholipids (34), respectively.
The formation of a mononuclear U(IV) phase has also been reported for other bacteria
of relevance to U bioremediation (7, 15), yet contrasts with earlier reports of uraninite
formation by Geobacter spp. (17, 62). The chemical composition of the medium can influence
the nature of the reduced U mineral (15). We used a bicarbonate buffer and conditions used in
previous studies with G. sulfurreducens (63), whereas studies reporting uraninite formation
used PIPES-buffered solutions (62) or bicarbonate-buffered uncontaminated groundwater (17).
49

Evidence for the microbial reduction of U(VI) to non-uraninite U(IV) products is also emerging
from field-scale studies (7, 29), whereas uraninite formation has been linked to conditions of
reduced bioreducing activities (27, 30). This suggests that abiotic factors may contribute to the
formation of uraninite.
Model for the reduction of U(VI) by Geobacter bacteria. The direct correspondence
observed between piliation, extent of U(VI) reduction, cell envelope respiratory activities and
cell viability support a model in which the conductive pili function as the primary mechanism
for U reduction and cellular protection (Fig. 2.9).
U(VI)
U(IV)

LPS
OM
PS
IM
PilA
FIGURE 2.9: Model for the extracellular reduction of U(VI) to U(IV). Conductive pili function as
primary uranium reductases, with c-cytochrome foci (c-cyt) as secondary reduction sites. IM,
inner membrane; PS, periplasmic space; OM, outer membrane; LPS, lipopolysaccharide; PilA,
pilin subunit.
Pili can reach several m in length, thereby increasing the redox-active surface area available
for binding and reducing U(VI) outside the cell. Although most of the U reduced by the piliated
50

cells was extracellular and associated with the pili, discrete regions of the outer membrane also
participated in the reduction of U. In G. sulfurreducens most of the redox activity of the outer
membrane is provided by abundant c-cytochromes that decorate the cell surface as defined
foci (52). Thus, they could provide a mechanism for reducing U in localized regions of the
membrane and preventing it from permeating into the periplasm. In support of this, the PilA

-

mutant cells, which had reduced outer membrane cytochrome content, preferentially reduced
U in the periplasm. Some of the most abundant metalloproteins on the outer membrane of G.
sulfurreducens also are loosely bound to and easily detach from the membrane (46, 47, 52),
providing a natural mechanism for releasing the U deposits. Some of these cytochromes also
may be anchored to a recently identified exopolysaccharide matrix (56), which may promote
the extracellular reduction of U. Furthermore, although some areas of the outer membrane
were devoid of U reductase activity, U(VI) was effectively prevented from permeating into the
periplasm. The outer leaflet of the outer membrane of Gram-negative bacteria is mostly
composed of lipopolysaccharide (LPS), which acts as an efficient permeability barrier against
soluble toxic compounds (53). G. sulfurreducens produces a rough LPS, i.e., it is composed of
lipid A and a core oligosaccharide but lacks the O-antigen (69). The core oligosaccharide is the
most highly charged region of the LPS and is stabilized by metallic cations (60). Models suggest
that rough LPS preferentially chelates and immobilizes uranyl ions over other ions (37) and
produces carboxyl and hydroxyl coordinations (37), which is consistent with the C and O ligands
modeled from the EXAFS spectra. This suggests that the rough LPS of G. sulfurreducens also
functions as a protective barrier to prevent U(VI) from penetrating in the cell envelope.

51

Implications for the in situ bioremediation of U. An insufficient knowledge of the
biological mechanisms of contaminant transformation often limits the performance of in situ
subsurface bioremediation and long-term stewardship strategies. The identification of
Geobacter’s pili as their primary U reductase provides a much-needed, fundamental
mechanistic understanding of U reduction by Geobacter spp. required to design effective in situ
bioremediation strategies. Analyses of transcript abundance for key Geobacteraceae genes are
useful tools to predict the metabolic and physiological state of Geobacter bacteria during in situ
bioremediation (21-24), yet provide no information about the mechanism of U bioreduction.
However, similar tools could be applied to monitor the activity of conserved components of
Geobacter’s pilus apparatus to assess the effectiveness of in situ bioremediation schemes. The
possibility that conductive appendages such as the pili of Geobacter are a widespread
mechanism for U reduction also warrants special attention. The production of conductive
appendages has been demonstrated in another U-reducing bacterium, Shewanella oneidensis
(18). Furthermore, nanowire-mediated electrical currents have been proposed to couple
spatially separated geochemical processes in sediments (50). The extracellular needle-like U
deposits observed in TEM thin sections of the piliated strains of G. sulfurreducens (Fig. 2.5) also
resembled the uraninite structures of Desulfovibrio desulfuricans biofilms (44), which some
authors have suggested represent mineralized nanowire-like appendages (71). Thus, the
contribution of microbial nanowires to U reduction may be significant and, therefore, relevant
for the optimization of in situ bioremediation strategies.

52

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60

CHAPTER 3

THE ROLE OF GEOBACTER SULFURREDUCENS BIOFILMS IN THE IMMOBILIZATION
AND REDUCTION OF URANIUM

The following individuals contributed to the work presented in this chapter:

Allison Speers: Analysis of COMSTAT data, collaborated on U experiments
Blair Bullard: Assisted with the development of biofilm and U assays
Shelly Kelly (EXAFS Analysis): EXAFS experiments and data analysis

61

ABSTRACT
Biofilms formed by dissimilatory metal reducers are of interest to develop permeable
biobarriers for the immobilization and reductive precipitation of soluble contaminants such as
uranium. Thus, we studied the kinetics of U(VI) removal and reduction by biofilms formed by
the metal-reducing bacterium Geobacter sulfurreducens. The biofilms removed twofold more
U(VI) from solution than planktonic cells, and linearly for up to 24 h. Despite the prolonged
exposure to uranium, the biofilm cells were viable and active, as indicated by their respiratory
activity, which was higher than in control biofilms not exposed to uranium. Furthermore, we
measured similar respiratory activities in cells from biofilms exposed to concentrations of
uranium as high as 2.5 mM, and approximately 70% of this activity was still detected in biofilm
cells exposed to 5mM concentrations for 24 h. Thus, the biofilms were able to maintain their
redox activities despite the high concentrations of uranium and prolonged exposure. The ability
to remove U(VI) from solution was similar in monolayered and multilayered biofilms and was
not proportional to increases in biofilm biomass and thickness. Hence, the ability to remove
uranium depended on the area of the biofilm that was exposed to the soluble contaminant. By
contrast, the reduction of uranium increased as the biofilms grew in biomass and height and
was dependent on the expression of Geobacter conductive pili. As with pili-expressing
planktonic cells, the reduced uranium mineral associated with the biofilms was a mononuclear
U(IV) phase involving carbon ligands. Taken together these results demonstrate that the stage
of biofilm development, as well as the specific components expressed at that stage determines
its contribution to U transformations.

62

INTRODUCTION
Since World War II uranium (U) has been mined extensively for its use in nuclear weapons,
and as a power source, resulting in the production of significant amounts of contaminated
groundwater and sediment worldwide (1, 18). One remediation strategy that has shown
promise is the stimulation of the native microbial community in situ. With the addition of an
electron donor, the native microbes are able to reduce soluble U(VI) to sparingly soluble U(IV),
thus preventing migration of the contaminant in groundwater (2, 9, 20, 31, 43). Concurrent
with the reduction of U(VI) is the enrichment the Geobacteraceae family of dissimilatory metalreducing organisms (2, 9, 31, 42).
The energy for growth of the Geobacteraceae during in situ bioremediation comes from the
reduction of Fe(III) oxides, an electron acceptor more prevalent than U in the subsurface (13).
The reduction of Fe(III) oxides requires the expression of Geobacter’s conductive pilus
nanowires (34), which recent studies from our group have also implicated as their primary U
reductase (10).
The expression of conductive pili by Geobacter also leads to cell aggregation and the
formation of biofilms (35, 36). Biofilm development is often assumed in the subsurface,
particularly at the matrix-well screen interface and rock fractures, but evidence of biofilms in
the bulk aquifer matrix is scarce (6). Recent studies (23) at the Rifle IFRC site demonstrated that
field-scale addition of acetate to groundwater also stimulated the growth of Geobacter spp. in
the sediment particles. Furthermore, their growth shifted from the groundwater to the solid
phases during the field-scale acetate addition, where they out-competed other organisms. This
suggested that Geobacter cells transitioned from planktonic to biofilm physiologies during the
63

active phase of U reduction following the addition of the electron donor. However, the
contribution of Geobacter biofilms to uranium removal and reduction has never been
investigated. This contrasts with the availability of studies about U transformations mediated
by Geobacter planktonic cells (10, 14, 25, 27, 40) or by biofilms formed by other metal-reducing
bacteria such as Desulfovibrio (5, 29) and Shewanella (7, 39).
Hence, we investigated the role of Geobacter biofilms in the immobilization and reduction of
uranium using the model representative Geobacter sulfurreducens, for which a genetic system
(11), sequenced genome (30), and developed biofilm protocols have been described previously
(35, 36). Here we show that G. sulfurreducens biofilms immobilize and reduce more uranium
than planktonic cells and are also able to tolerate exposure to higher concentrations of the
contaminant for prolonged periods of time. As with planktonic cells, the biofilms reduced the
soluble U(VI) to a mononuclear U(IV) mineral phase that included carbon ligands. The ability to
immobilize uranium was independent of the biofilm biomass and height. However, U reduction
correlated well with increases in biofilm biomass and height and was dependent on the
expression of the cell’s conductive pili. These findings support the notion that Geobacter
biofilms contribute to the immobilization and reduction of U in the subsurface, and highlight
their potential as permeable biobarriers for the bioremediation of uranium contaminants.

MATERIALS AND METHODS
Strains and culture conditions. Wild-type (WT) G. sulfurreducens PCA (ATCC 51573), a
-

pilin-deficient mutant (PilA ) (34), and its genetically complemented strain (pRG5::pilA) (34)
were routinely cultured in fresh water (FW) medium (26) with the modifications described
64

previously (10), and supplemented with 15mM acetate and 40mM fumarate (FWAF). The
medium was dispensed into serum bottles, sparged with N2:CO2 (80:20), sealed with butyl
rubber stoppers (Bellco) and aluminum tear-off seals (Wheaton), and autoclaved 30 minutes.
Biofilms were grown on 6-well cell-culture-treated plates (Corning), or glass coverslips (Corning)
(36). Prior to inoculation the glass coverslips were acid-washed overnight with a 50/50 (vol/vol)
-

HCl/NO3 mixture or a 15% (vol/vol) HCl/H2O mixture, rinsed thoroughly with ddH2O, and
inserted into sliced rubber stoppers (4 coverslips/stopper), as previously described (36).
Immediately prior to inoculation, the stopper-coverslip assembly was autoclaved in FW medium
lacking vitamins, minerals, acetate, and fumarate. Each stopper-coverslip assembly was placed
in a sterile 50 ml conical tube (Corning). Biofilm assays were inoculated with an early stationaryphase FWAF culture to a final OD600 of 0.04, grown anaerobically inside a vinyl glove bag (Coy
Labs) with a H2:CO2:N2 (7:10:83) atmosphere, and incubated at 30°C for 24, 48, or 72 h, as
specified.
For determination of the total protein content of the biofilms, biofilms were grown for
24, 48 or 72 h in 6-well plates, scraped off, and harvested by centrifugation (5 min, 12,000 x g).
The resulting cell pellet was boiled for 1 h in 2M NaOH, allowed to cool, and then neutralized
with an equal volume of 2M HCl. The sample was centrifuged to remove cellular debris, and the
resulting supernatant analyzed for total protein content. Protein was quantified using a Pierce
Microplate BCA Protein Assay Kit (reducing reagent compatible, Thermo Scientific) with BSA

65

standards, according to the manufacturer’s specifications. Protein was measured as an OD562
on a Tecan Sunrise Plate Reader (Tecan, Inc.).
U(VI) resting cell suspension and biofilm assays. The ability of cells to remove U(VI),
provided as uranyl acetate, from solution was assayed in resting planktonic and biofilm cell
suspensions using protocols adapted from those described previously (40). Heat-killed and
uninoculated controls were also included to rule out any abiotic removal activity or absorption.
Resting biofilm suspensions were prepared from biofilms grown on stopper-coverslip
assemblies for 24, 48, or 72 h, as described above. The culture broth was decanted, the
assembly rinsed gently with sterile, anaerobically-prepared wash buffer (40), and 20 ml of
reaction buffer (40) supplemented with 20 mM sodium acetate and 1 mM uranyl acetate
(Electron Microscopy Sciences) prepared in 30mM bicarbonate buffer was added. Resting
planktonic and biofilm cell suspensions were incubated for 24 h at 30°C. Following incubation,
500 l samples of the supernatant were removed, filtered (0.22 µm Millex-GS filter, Millipore),
acidified in 2% nitric acid (500 L), and stored at -20°C. For kinetic studies of U(VI) removal,
samples were taken every 6 h. All procedures were performed inside an anaerobic glove bag, as
described above. The concentration of U(VI) in the acidified samples was measured using a
Platform Inductively Coupled Plasma Mass Spectrometer (ICP-MS) (Micromass,Thermo
Scientific) or a Kinetic Phosphorescence Analyzer (KPA) (Chemchek).
Vitality fluorescent assays. The respiratory activity of biofilm cells after exposure to
uranium was assayed using the fluorescent RedoxSensor vitality kit (Invitrogen), as previously
described. WT biofilms were grown on coverslips for 48 h and incubated in reaction buffer with
66

1 mM, 2.5 mM and 5 mM concentrations of uranyl acetate. Control biofilms incubated in
reaction buffer without uranyl acetate were also included. After 24 h of incubation, the reaction
buffer was decanted from the tubes and the stopper-coverslip assemblies were washed with
wash buffer. The biofilms were then scraped from the assembly and resuspended in 1 ml
reaction buffer. Samples were vortexed briefly, mixed 1:1 with Redox dye solution, and
incubated 10 min before measuring fluorescence (490nm excitation, 520nm emission) on a
SpectraMax M5 plate reader (Molecular Devices). The respiratory activity of the biofilms was
calculated as the fluorescence emission of the Redox Sensor dye relative to the metabolic
activity of controls without uranium. Separate aliquots of the samples were stained with SYTO 9
(Invitrogen) to confirm that the samples had the same amount of cells.
Microscopy. Biofilm growth on 6-well plates was examined by Confocal Laser Scanning
Microscopy (CLSM). Following the specified incubation period, planktonic growth was carefully
removed and the remaining biofilm was stained with LIVE/DEAD BacLight Bacterial Viability Kit
(Invitrogen) dye solution, following the manufacturer’s recommendations. The biofilms were
stained for approximately 15 min, washed once in PBS, and imaged on a Zeiss Pascal LSM
microscope (Carl Zeiss Microscopy, LLC) equipped with an Achroplan 40x/0,80W dipping
objective. COMSTAT analyses were carried out using images from three biological replicates,
with 6-10 distinct fields-of-view (1,024 x 1,024 pixels, 0.22 µm/pixel) for each. Images were
collected every 1.14 µm, and projections were created using Zeiss LSM Image Browser software
(Carl Zeiss Microscopy, LLC). The structure of the biofilms was characterized using COMSTAT
image analysis software using connected volume filtration to remove noise in the data, as
described previously (17).
67

When indicated, the biofilms were also examined with a Scanning Electron Microscope
(SEM). Biofilms were grown for 48 h on round glass coverslips (12-mm diameter), and exposed
to 1mM uranyl acetate for 24 h. The biofilms were then fixed at 4°C for 1-2 h in 4%
glutaraldehyde, rinsed briefly in 0.1M sodium phosphate buffer, and dehydrated in a series of
ethanol washes (25%, 50%, 75%, 95%, 10 minutes each), followed by three 10 min washes in
100% ethanol. The samples were critical-point dried using a Blazers 010 critical point dryer
(Blazers Union Ltd.) with liquid CO2 as the transitional fluid. Once dry, the coverslips containing
the biofilm samples were mounted on aluminum stubs using epoxy glue and coated with ~10
nm of osmium using a NEOC-AT osmium coater (Meiwafosis Co., Ltd.). Samples were examined
with a JEOL JSM-7500F SEM equipped with an Energy Dispersive Spectroscopy (EDS) 30mm

2

detector crystal for elemental analyses.
X-ray Absorption Spectroscopy (XAS) analyses. The valence and speciation of U in the
biofilms was estimated by XANES (X-ray Absorption Near Edge Spectroscopy) and EXAFS
(Extended X-ray Absoption Fine Structure Analysis), respectively. For these analyses, biofilms
were grown on stopper-coverslip assemblies, and exposed to U for 24 h, as described above.
After rinsing the assemblies gently with wash buffer, the biofilm biomass was scraped off the
assemblies using and resuspended in 2 ml of reaction buffer. The cells were then harvested by
centrifugation (12,000xg, 10 min), loaded into custom-made plastic holders, and stored at 80°C (10). All procedures were carried out in an anaerobic chamber, and samples were kept
frozen during XAS measurements. XANES and EXAFS measurements were performed using
standard beamline parameters (22) and a multielement Ge detector in fluorescence mode using
68

the PNC-CAT beamline 20-BM at the Advanced Photon Source (Argonne National Laboratory).
Data obtained from XANES and EXAFS analysis was processed according to the combination of
protocols described previously (10).
SDS-PAGE and heme-stain of proteins of the biofilm EPS matrix. The exopolysaccharide
matrix (EPS) was extracted from biofilms grown 48 h on 6-well plates using a modification of a
protocol described previously (8, 38). Briefly, biofilms were scraped and collected in reaction
buffer. The solution was centrifuged for 10 minutes at 13,000xg and the resulting pellet was
resuspended in 1/5 volume of TNE and vortexed for 1 min. SDS was added to a final
concentration of 0.1%, and the solution was mixed at room temperature for 5 min. The samples
were then passed 10 times through an 18G needle, and centrifuged at 15,500xg to collect the
sheared materials as an insoluble fraction. The resulting pellet was washed 5 times before
resuspending it in 10mM Tris-HCl, pH 7.5.
To identify heme-containing proteins in the EPS matrix, 20 µg of protein from each EPS
isolation was boiled for 10 min and separated on a 12% Mini-Protean TGX gel (BioRad) at 250V
for 30 min. The Novex Sharp markers (Invitrogen) were used a molecular weight standard.
Heme-containing proteins were visualized on the gel with N,N,N’,N’-tetramethylbenzidine
staining, as described previously (10, 41). A duplicate gel was run in parallel and stained for
total protein using Coomassie Brilliant Blue G-250 (BioRad) according to the manufacturer’s
recommendations.

69

RESULTS AND DISCUSSION
Enhanced U(VI) immobilization and tolerance by biofilms. The kinetics of U(VI)
immobilization were investigated in resting 48 h biofilms in reference to planktonic cells (Fig.
3.1A).

FIGURE 3.1: Biofilms maintain vital activities through 24 h of U exposure. (A) Removal of U(VI)
from solution by 48 h biofilms of G. sulfurreducens (solid symbols) showing the linearity of the
reaction for 24 hours. Controls with planktonic cells (open symbols) and uninoculated biofilm
assemblies (dashed line) are also shown. (B) Effect of 24 h exposure to increasing
concentrations of U(VI) (1, 2.5, and 5 mM) on the respiratory activity of 48-h biofilms (vitality
measured with the RedoxSensor dye and expressed as the ratio of relative fluorescence units
+

−

emitted by cells incubated with (U ) and without (U ) U).

The rates of U removal during the first 6-12 h were similar in planktonic cells and
biofilms. However, while the biofilms immobilized U linearly for up to 24 h, the removal
activities of the planktonic cells stopped after 12 h and the cell-associated U was solubilized
again. As a result, the biofilms immobilized twice as much U as the planktonic cells. Cell
viability, which is crucial for assessing the capacity for U(VI) reduction, can only be preserved in
70

planktonic cells for the first 7 h under the conditions used in this assay (40). After this time,
osmotic pressure causes the cells to lyse and the viability of the resting cells declines rapidly.
The sustained removal of U by the biofilms suggests that the biofilms remain viable and
metabolically active even after prolonged exposure to the contaminant. To investigate this, we
used the fluorogenic RedoxSensor Green dye to measure the respiratory activities of biofilm
+

-

cells exposed to U for 24 h (U ) in reference to unexposed biofilm controls (U ) (Fig 3.1B). The
dye yields green fluorescence when modified by the bacterial reductases, which are mostly
located in the electron transport system of the cell envelope (15, 21). As respiration is a vital
activity of the cell, the dye also measures the cell’s vitality and serves as a proxy for the cell’s
viability (10). As shown in Fig. 3.1B, the respiratory activities of biofilm cells exposed to 1mM U
for 24 h were 2.4-fold higher than control biofilms incubated under identical conditions but
without U. This is likely because they are able to use the U as an electron acceptor, while the
unchallenged controls are not supplemented with an electron acceptor and therefore are not
actively metabolizing. This high rate of vitality contrasts with the more than 70% decrease in
respiratory activities reported for planktonic cells incubated with U for 6 h in reference to
unexposed controls (10). Furthermore, similar increases in respiratory activities were measured
in biofilms exposed to 2.5 mM concentrations of U for 24 h and decreases in respiratory activity
(ca. 70%) were only measured after exposing the biofilms to 5 mM concentrations of U for the
same period of time. Thus, the results are consistent with increased respiratory activities and
cell viability in cells within biofilms.
Extracellular reduction of U by biofilms. Approximately 67% (±4%) of the U(VI)
immobilized by 48 h biofilms after 24 h was reduced to U(IV). Thus, the immobilization of U(VI)
71

by the biofilms was coupled to its reduction of U(IV). Examination of the biofilms after exposure
to 1 mM U for 24 h by SEM showed needle-like, extracellular precipitates that coated the
biofilm microcolonies (Fig. 3.2A), which elemental analyses with an EDS detector confirmed to
be composed of U (Fig. 3.A.1).

FIGURE 3.2: SEM micrographs of 48 h biofilms exposed to U for 24 h (A and B) showing the
extracellular needle-like, white precipitates of uranium associated with the biofilm
microcolonies. Control biofilms not exposed to U are also shown (C). Scale bar, 1 µm.

At higher magnification (Fig. 3.2B and C) the U precipitates were observed as
interspersed with extracellular filaments, some with diameters (ca. 4 nm) matching well the
diameters reported for the conductive pili of G. sulfurreducens (34) and some with larger
diameters (ca. 15-20 nm) consistent with dehydrated EPS fibers (38). The conductive pili of G.
sulfurreducens are required for microcolony formation during biofilm formation by G.
sulfurreducens (36) and are also the primary U reductase of planktonic cells (10). Thus, they
could also play a similar role in U reduction by the biofilms. On the other hand, the EPS matrix
of G. sulfurreducens anchors several c-cytochromes involved in metal reduction in G.

72

sulfurreducens (38) and could, therefore, contribute to the U reductase activities of the
biofilms.
As the biofilms grow and mature, more conductive pili and more EPS matrix with ccytochromes are available to participate in the redox reactions of the biofilms. Hence, we
studied the ability of biofilms grown for 24, 48 and 72 h to immobilize and reduce U(VI) after 24
h of exposure to the contaminant (Fig. 3.3A).

-

FIGURE 3.3: U(VI) immobilization/reduction, and cytochrome profile of the WT, PilA , and
pRG5::pilA strains. (A-B) Reduction of U(VI) (open blocks) to U(IV) (solid blocks) by WT biofilms
-

grown for 24, 48 and 72 h (A) and by 48 h biofilms of the pilin-deficient PilA mutant and the
hyperpiliated pRG5::pilA strain (B). (C) Heme-stained SDS-PAGE of protein extracted from the
-

EPS matrix of 48 h biofilms of the WT (lane 1), PilA (lane 2), and pRG5::pilA (lane 3) strains.
Numbers at the right are relative molecular weights of protein markers in kDa. The arrow points
at the migration of the small, processed form of the OmcZ c-cytochrome (OmcZS).

While the removal activities of the biofilms were similar, U reduction increased
proportionally to the biofilm age and was highest in the 72 h biofilms (Fig. 3.3A). CLSM
micrographs of the biofilms revealed visual differences in biofilm thickness and structure in 24,
48, and 72 h biofilms (Fig. 3.4A and 3.A.2).
73

FIGURE 3.4: CLSM micrographs showing side view projections of (A) WT (24, 48 and 72 h), (B)
-

PilA (48 h), and (C) pRG5::pilA (48 h) biofilms stained with the BacLight viability kit (green,
live cells; red, dead cells). Scale bar, 20 µm. The top view projections corresponding to these
images are shown in Fig. 3.A.2.

Biofilm parameters calculated using the COMSTAT analysis software such as biomass,
thickness and surface area, increased linearly during the first 48 h of biofilm growth and
remained unchanged in 72 h biofilms (Fig. 3.A.3, Table 3.1). Interestingly, the surface coverage
and surface to volume ratio values did not change substantially as the biofilms aged (Fig. 3.A.3,
Table 3.1).

74

TABLE 3.1: COMSTAT analysis
Biofilm

Total biomass
3

2

(µm /µm )

Average
Thickness
(µm)

Surface
Area

Surface to volume

(mm )

Surface
Coverage
(%)

2

2

3

ratio (µm /µm )

WT 24h

6.4 ± 1.0

7.8 ± 1.1

0.6 ± 0.1

80.7 ± 5.2

2.9 ± 0.3

WT 48h

10.6 ± 1.3

13.3 ± 1.8

1.2 ± 0.1

91.8 ± 1.7

3.6 ± 0.4

WT 72h

10.6 ± 3.3

13.9 ± 3.9

1.0 ± 0.2

92.2 ± 6.6

3.1 ± 0.4

PilA 48h

-

5.2 ± 0.7

6.8 ± 1.0

0.6 ± 0.1

77.8 ± 3.6

3.4 ± 0.3

pRG5::pilA
48h

43.4 ± 14.8

60.1 ± 18.9

6.1 ± 2.5

99.5 ± 0.5

4.4 ± 1.3

This is similar to our earlier observation that the U removal activities of the biofilms do
not change substantially as the biofilms age (Fig. 3.3A). This and the extracellular location of the
U precipitates (Fig. 3.2) suggest that U is preferentially immobilized at the biofilm surface
exposed to liquid milieu.
None of the biofilm parameters calculated (Fig. 3.A.3) matched the linear increases in U
reductive activities observed as the biofilms aged between 48 and 72 h (Fig. 3.3A). Biofilm
formation is a developmental process, consisting of specific stages such as attachment,
microcolony formation and biofilm maturation (32). The transition from one stage of biofilm
development to the next involves extensive gene reprogramming so that specific biofilm
components and activities are expressed (16, 28). Consistent with this, we measured linear
increases in the total protein content of the biofilms, which includes the protein from the cells
75

and from the biofilm matrix (Fig. 3.A.4), that cannot be accounted for by increases in cell
numbers calculated as biomass in the COMSTAT analyses (Fig. 3.A.3). This supports the notion
that there are specific redox-active components expressed during biofilm formation that
catalyze the reduction of U.
Role of conductive pili and c-cytochromes of the biofilm matrix in U reduction. We
further investigated the role of the biofilm pili in U reduction by studying the ability of 48 h
-

biofilms of a pilin-deficient (PilA ) mutant to immobilize and reduce U compared to its
genetically complemented strain pRG5::pilA (Fig. 3.3B). The mutant carries a deletion in the
gene encoding the pilin subunit (34), which reduces the ability of planktonic cells to reduce U
extracellularly (10). The mutant biofilms attached and grew on the surface but had fewer
microcolonies (Fig. 3.4B and Fig. 3.A.2). By contrast, it’s genetically complemented strain,
pRG5::pilA, which is hyperpiliated (10), formed very thick biofilms after only 48 h of incubation
(Fig. 3.4C and Fig. 3.A.2). In general, all the biofilm parameters measured in the mutant biofilms
(such as biomass, thickness, surface coverage, etc.) were similar to those measured in 24 h
biofilms of the WT strain (Fig. 3.A.3), consistent with the previously reported role of Geobacter
pili in microcolony formation (36). Not surprisingly, the mutant biofilms also removed and
reduced less U than 48 h WT biofilms (Fig. 3.3B). By contrast, the hyperpiliation of the strain
pRG5::pilA promoted biofilm formation (Fig. 3.4C and 3.A.2) and resulted in substantial
increases in biofilm biomass, thickness and surface area but similar values for surface coverage
and surface:volume ratio (Fig. 3.A.3). The hyperpiliated biofilms also removed 1.6 times more U
than the WT biofilms and reduced 76% (±3%) of U(VI) to U(IV) (Fig. 3.3B).
When compared to 48 h WT biofilms, the U removal and reduction activities of the PilA
76

-

and pRG5::pilA biofilms correlated strongly with the biofilm biomass and average thickness (R

2

= 0.99), but required a logarithmic fit. This indicates that a biofilm’s ability to adsorb or reduce
U based solely on its biomass characteristics is finite. Hence, increases in biofilm biomass
cannot solely account for the differences in U removal and reduction observed in the biofilms
formed by strains with different levels of piliation. Interestingly, we observed a positive linear
2

correlation between surface coverage and U removal (R = 0.82). Surface coverage represents
the amount of cells attached to the substratum, or biofilm confluence (17). The roughness
coefficient is a measurement of the variations in biofilm thickness (17). Thus, a lower roughness
coefficient indicates a more uniform biofilm and a higher coefficient a more variable one. Taken
together these results demonstrate that the thickness/biomass of a biofilm can be used as an
initial prediction of the amount of U that can be immobilized and reduced, but the ultimate
determination of the U transformation phenotype is made by structural characteristics
determined by specific biofilm components, such as the pilus nanowires.
-

As the PilA mutant also has defects in outer membrane c-cytochromes required for
metal reduction such as OmcS (10) and these cytochromes are anchored in the biofilm EPS
matrix (37), we examined the heme-containing proteins associated with the biofilm matrix of
-

the WT, PilA and pRG5::pilA biofilms (Fig. 3.3C). Interestingly, a band with a relative molecular
-

weight similar to that of OmcS (~47KDa) (33) was present in both the WT and the PilA mutant
matrices. The only heme-containing band present in the WT matrix and absent in the PilA

-

matrix was one with a relative molecular weight of 30 KDa. This size matches that of OmcZS, a
77

processed isoform of the OmcZ cytochrome of G. sulfurreducens that is required for optimal
current production by anode biofilms in microbial fuel cells (19). In vitro studies show that the
purified OmcZS protein can reduce U (19). Thus, OmcZS could contribute to the reduction of U
in the biofilms as well. Interestingly, the pRG5::pilA heme-stain profile also shows defects in
OmcZS levels (Fig. 3.3C), yet produces more conductive pili and reduces more U than the WT
2

biofilms (Fig. 3.3B). Furthermore, we observed a strong linear correlation (R = 0.999) between
the levels of piliation of the three strains grown under pili-inducing conditions (25°C) and the
extent of U reduction by the biofilms (Fig. 3.A.4B). Hence, the results support the notion that
the conductive pili are the primary U reductase of the biofilms.
-

XAS analyses. U L3-edge EXAFS spectra from WT and PilA biofilms were modeled to
determine the atomic coordination about U. The spectra were best described by a mixture of
U(IV) and U(VI) coordinated by carbon atoms. The magnitude of the Fourier transformed
spectra and models are shown in Fig 3.5A, with the spectra offset for clarity.

78

FIGURE 3.5: U LIII-edge XAFS spectra. EXAFS spectra and model. (A) Magnitude of Fourier
-

transform spectra are offset for clarity in descending order PilA , and WT. The real part of
-

Fourier transform of WT (B) and PilA (C) is also shown. The components of the model are
shown offset beneath the total model and measured spectrum. Also shown is the U(IV) moiety
-

that is consistent with the measured EXAFS spectra for both WT and PilA (inset). U(IV) (grey) is
coordinated by oxygen (red) atoms and two bidentate carbon (black) ligands and one
monodentate carbon ligand.

79

-

A molecular moiety that is consistent with both the WT and PilA model is shown in the
inset. Fig 3.5B and 3.5C show the contribution of each path in the model in the real part of the
-

Fourier transform for the WT and PilA samples, respectively. The model includes two types of
carbon ligands. One of the C-ligands is bound to two oxygen atoms of U in a bidentate fashion
and is followed by a distant carbon (C3) atom. The other C-ligand is bound to one oxygen atom
of U in a monodentate fashion and is attached to 3 distant oxygen (Odist) atoms. Multiple
scattering paths from distant C3 and Odist atoms were included in the model. This model was
2

simultaneously refined to both spectra. The distances and σ values used to model the spectra
are listed in Table 3.A.1. Table 3.A.2 lists the coordination numbers. The coordination numbers
are consistent with 1 to 2 bidentate C-ligands and 1 to 2 monodentate C-ligands per U atom.
The number of Oax atoms can be used to estimate the amount of U(IV) in these samples, since
there are two Oax atoms for each U(VI) atom and no Oax atoms for U(IV). The Noax values
-

indicate that PilA and WT contain approximately 25%, and 48% U(IV) with an estimated
uncertainty of 10%. These values are consistent with U XANES measurements indicating more
-

U(IV) in the WT sample as compared to the PilA sample.
Our previous studies with planktonic cultures required the addition of a phosphorus
-

ligand to model U reduction in the PilA strain (10), which we hypothesized was the result of U
permeating deep into the cell envelope and complexing with phosphorous-containing
substrates such as peptidoglycan (3), periplasmic proteins (4), and membrane phospholipids
-

(24). Interestingly, although the PilA biofilms showed a distinct U removal and reduction
80

defect, the model that fit the EXAFS spectra did not require a phosphorous ligand, suggesting
that U may not have permeated in the cells but, rather, was immobilized and reduced by other
extracellular components of the biofilm matrix. Biofilm physiology is substantially different
from that of planktonic cultures (12). Thus, other yet to be identified biofilm components are
likely to contribute to the redox activities of the biofilms.
Implications for bioremediation. The results of this study demonstrate the relevance of
biofilms to U transformations in the subsurface, and highlight the distinct contributions of each
stage of development, as well as the necessity to better understand the physiology of bacteria
in the subsurface. Further studies into the U reduction activity and development of Geobacter
biofilms could contribute to the implementation of novel bioremediation strategies, potentially
including the use of biofilms as bioreactive barriers.

81

APPENDIX

82

FIGURE 3.A.1: SEM micrograph and EDS analysis of U precipitates. (A) SEM micrograph of 48 h
biofilms exposed to U for 24 h, with the area of EDS analysis indicated by the red box. (B) EDS
spectrum of the white precipitates within the boxed region, with U peaks indicated by red
arrows. Scale bar, 1 µm. The text in this figure is not meant to be readable, but is for visual
reference only.

83

FIGURE 3.A.2: CLSM micrographs showing top view projections of 24, 48, and 72 h WT (A), 48
-

h PilA (B), and 48h pRG5::pilA (C) biofilms stained with BacLight viability kit (green, live cells;
red, dead cells). Scale bar, 20 µm.

84

FIGURE 3.A.3: Correlation of biofilm age and biofilm biomass (A), thickness (B), surface area
(C), surface coverage (D), and surface:volume ratio (E). Values were determined using
-

COMSTAT analysis software (17). WT (black circles), PilA (squares), and pRG5::pilA (triangles)
strains are shown.
85

FIGURE 3.A.4: Linear correlation between the total protein content of biofilms and biofilm
2

age (R =0.94) (A) and between piliation (planktonic cells grown under pili-inducing conditions
-

2

at 25°C) and U reduction by 48 h biofilms of the WT, PilA and pRG5::pilA strains (R = 0.999)
(B).

86

2

Table 3.A.1: EXAFS modeling results for R and σ
R (Å)

**

σ2 (∙10

-3 2

Path

CN

Oax

Noax

1.80 ± 0.01

1*

Oeq
C1
C2

2.38 ± 0.02
2.86 ± 0.01
3.49 ± 0.03

**
2±7
2±7

Oax1-Oax2

Noeq
Nc1
Nc2
Noax

3.61 ± 0.02

2*

Oax1-U-Oax2

Noax

3.61 ± 0.02

2*

Oax1-U-Oax1
C3
C1-C3
C1-C3-C1
Odist
C2-Odist
C2-Odist-C2

2Noax
Nc1
2Nc1
Nc1
Nc2
2Nc2
Nc2

3.61 ± 0.02
4.54 ± 0.07
4.54 ± 0.07
4.54 ± 0.07
4.69 ± 0.08
4.72 ± 0.08
4.74 ± 0.08

4*
2±7
2±7
2±7
2±7
2±7
2±7

Å )

*value held, **PilA: 25 ± 6, WT: 18 ± 4

Table 3.A.2: EXAFS modeling results for coordination
numbers
Data Set
Noax
Noeq
C1
C2
PilA1.5 ± 0.1
6.9 ± 2.2
1.2 ± 0.7
1.2 ± 0.7
WT
1.0 ± 0.1
6.3 ± 1.4
1.4 ± 0.7
1.4 ± 0.8

87

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88

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92

CHAPTER 4

GENETIC ANALYSIS OF BIOFILM DEVELOPMENT IN GEOBACTER
SULFURREDUCENS

The following individuals contributed to the work presented in this chapter:

Allison Speers:
Collaborated on the development of a mutagenesis system, and the optimization of the biofilm
assay used for the mutant screening.
Anne Otwell and John Rotondo:
Undergraduate researchers who assisted with the construction of the mutant library, biofilm
screening assays, and sequencing associated with this project.

93

ABSTRACT
As discussed in Chapter 3, Geobacter sulfurreducens biofilms are able to reduce uranium
for prolonged periods of time without compromising the cell’s viability. Our studies also
demonstrated that each biofilm developmental stage contributes differently to U
transformations, with reductase activity becoming more prominent in the later stages, and
independent of the quantity of biomass. This led us to investigate the components expressed at
each stage of growth in order to develop molecular markers that could allow us to assess the
developmental stage of Geobacter bacteria in the subsurface, and to better predict their U
reductase activity.
To identify genes involved in biofilm development, we developed a transposon
mutagenesis system coupled with a high-throughput biofilm screening assay. Our partial
screening of a transposon-insertion library allowed the identification of genes required for
attachment, microcolony formation, and biofilm maturation in G. sulfurreducens. Our results
confirmed the essential role of the pilus nanowires in biofilm development, and identified
additional genes with cell envelope biogenesis and electron transfer functions that have not
previously been implicated in biofilm formation by G. sulfurreducens.

94

INTRODUCTION
It is now widely accepted that, in their natural environment, bacteria live predominately
as part of surface-attached communities termed biofilms (14, 15, 17). For this reason, biofilm
development has often been assumed in the subsurface, particularly at the matrix-well screen
interface and rock fractures, but evidence of biofilms in the bulk aquifer matrix is scarce (7).
Dissimilatory iron reducers in the genus Geobacter are natural inhabitants of subsurface
aquifers and sediments (10). These bacteria gain energy for growth by coupling the oxidation of
electron donors such as acetate to soluble and insoluble electron acceptors (30). The natural
electron acceptor of Geobacter bacteria is insoluble Fe(III) oxides, which are abundant in
subsurface groundwater and sediments (19). In vitro studies with the model representative
Geobacter sulfurreducens have shown that the bacterial cells attach and grow as biofilms on
Fe(III) coatings, a process that requires the expression of G. sulfurreducens conductive pili to
provide structural support for the biofilms and mediate long-range electron transfer across the
multilayered community (47). G sulfurreducens also forms electroactive biofilms on the anode
electrode of a microbial fuel cell (46). Furthermore, the growth of the electroactive biofilm
requires the expression of the conductive pili and is proportional to current production (46).
Evidence is also emerging from environmental studies that Geobacter bacteria form biofilms in
the subsurface and that these biofilms mediate redox reactions such as the reductive
precipitation of soluble uranium contaminants. The field-scale addition of acetate to
groundwater stimulates planktonic growth and uranium reduction by Geobacter spp. (3).
However, the growth of the planktonic population shifted from the groundwater to the solid
phases during the field-scale acetate addition, and, once in the sediment, the Geobacter spp.
95

out-competed other organisms (25) . This suggested that Geobacter cells transitioned from
planktonic to biofilm physiologies during the active phase of U reduction following the addition
of the electron donor.
As shown in Chapter 3, we have demonstrated that the biofilms formed by G.
sulfurreducens can reductively precipitate uranium. The biofilms immobilized more uranium
than planktonic cells for prolonged periods of time without compromising the cells viability
even when exposed at high (up to 2.5 mM) concentrations of uranium, thus showing great
potential for the development of permeable biobarriers that immobilize uranium and limit its
mobility in the aqueous phases. The reduced uranium was a mononuclear phase and was
predominantly extracellular and associated with the biofilm pili and exopolysaccharide (EPS)
matrix. We observed a linear correlation between piliation and the uranium reductase activities
of the biofilms, suggesting that the conductive pili, as in planktonic cells (11), are the primary
site for uranium reduction in the biofilms. We also identified the extracellular c-cytochrome
OmcZS (22) in the EPS matrix and evidence suggested that it could contribute to uranium
reduction in the biofilms. However, the uranium reductase abilities of biofilms of a mutant
deficient in pili production and OmcZS expression were not completely abolished. Thus, other
biofilms components may contribute to the reduction of uranium by the biofilms.
Also in Chapter 3, we showed that the uranium reductase activities of G. sulfurreducens
biofilms increase as the biofilms mature, although the biofilm biomass did not change
substantially. Bacterial biofilm formation is a developmental process that goes through defined
stages as cells transition from planktonic growth to a sessile community. These stages consist of

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attachment to the surface, the formation of microcolonies, biofilm maturation, and dispersal
(39). Each developmental stage involves the expression of a unique set of genes, which confers
on the biofilm a distinct physiology. As biofilms grow and mature, for example, they display
increased resistance to antimicrobial agents and heavy metals (21, 32). Thus, it is important to
develop molecular markers that can allow us to assess the developmental stage of Geobacter
bacteria in the subsurface and predict the redox reactions that they can catalyze.
Transposon mutagenesis has been successfully used to identify genes involved in biofilm
development in a variety of organisms (31, 40, 41, 44, 48, 60). Transposon-insertion mutants in
genes required to transition from one developmental stage to the next are interrupted at that
specific stage and can be identified in high-throughput screenings that quantify the biofilm
biomass. Crystal violet, for example, is most often used to stain the biofilm biomass and the
biofilm-associated dye can then be solubilized with acids or organic solutions and quantified
spectrophotometrically (42). Transposon mutagenesis has also recently been used to identify
genes required for attachment and electron transfer to electrodes and Fe(III) oxides by G.
sulfurreducens (48, 49). This, and the availability of assays to investigate biofilm formation in
this organism (47, 49), prompted us to develop a transposon mutagenesis method and a highthroughput biofilm screening assay to investigate the molecular basis of biofilm development in
G. sulfurreducens. Here we show results from a partial screening of a transposon-mutant library
of G. sulfurreducens and the identification of genes required for attachment, microcolony
formation and biofilm maturation in this organism. The genetic screening confirmed the
essential role of the pilus nanowires in biofilm development in this organism, and also identified

97

a number of genes involved in cell envelope biogenesis and electron transfer that have not
been previously implicated in biofilm formation.

MATERIALS AND METHODS
Strains and culture conditions. Wild-type (WT) Geobacter sulfurreducens PCA (ATCC
51573) was routinely cultured at 30°C in NB medium supplemented with 15mM acetate and
40mM fumarate (NBAF) (12). When indicated, fresh water medium (11, 30) supplemented with
15mM acetate and 40mM fumarate (FW 15A/40F), fresh water medium supplemented with
30mM acetate and 40mM fumarate (FW 30A/40F), or minimal medium (DB) supplemented
with 30mM acetate and 40mM fumarate (DB 30A/40F) (34, 48, 53) were used. All media were
sparged with N2:CO2 (80:20), sealed with butyl rubber stoppers, and autoclaved prior to use. All
procedures were performed anaerobically inside a vinyl glove bag (Coy Labs, Grass Lake, MI)
containing a H2:CO2:N2 (7:10:83) atmosphere.
Biofilm assays. To determine the conditions most conducive to biofilm growth and highthroughput screening, a variety of growth media, commercially-available 96-well plates, and
chemical coatings with different surface characteristics (Table 4.A.1) were compared. Biofilms
were incubated at 30°C for 24, 48, or 72 hours, as indicated. After the specified incubation
period, planktonic growth was discarded and the biofilms were stained with 0.1% (w/v) crystal
violet for 20 min, as described previously (31, 42).
Four solvents were tested to solubilize the biofilm-associated dye (Table 4.A.1), and
their optimal absorption wavelength was also investigated. Each plate was de-stained for 20
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minutes with the appropriate solvent, and the optical density (OD) was measured at three
wavelengths contained by its maximum absorption peak (580 nm, 590 nm, and 600 nm). Unless
otherwise indicated, plates were de-stained with the best solvent (33% acetic acid) and the OD
of the crystal violet-acetic acid solution was measured at 580 nm using a SpectraMax M5 Plate
Reader (Molecular Devices).
Confocal Laser Scanning Microscopy (CLSM). Biofilms were cultured and imaged in flatbottomed Costar cell culture cluster 6-well plates (Corning), which have the same binding
properties as the Corning 96-well plates selected for the biofilm screening (Corning microwell
selection guide, 2011). Cells were grown as specified in FW 30A/40F, FW 15A/40F, or DB
30A/40F for 24, 48, for 72 h before discarding the liquid culture containing the planktonic cells.
The biofilm was stained approximately 15 min with the BacLight LIVE/DEAD fluorescent stain
(Invitrogen) according to the manufacturer’s recommendations, and the dye solution was
replaced with PBS for imaging. Biofilms were imaged using the 488 nm and 543 nm lasers on a
Zeiss LSM Pascal microscope (Carl Zeiss Microscopy) equipped with an Achroplan 40x/0,80W
dipping objective.
Transposon mutagenesis. Mid-log phase WT cultures were serially transferred at least
three times in NBAF medium and the cells were then harvested and treated to make
electrocompetent cells as described previously (12). The commercially available linear EZ-Tn5
Transposome complex (Epicentre Biotechnologies) (20) was electroporated into wild-type G.
sulfurreducens electrocompetent cells using an Eppendorf 2510 Electroporator operated at
1470 volts. Cells were recovered for ~18 h in NBAF medium and plated on NBAF plating media
50

supplemented with 50 µg/ml kanamycin (KM ) to select for insertion mutants. A library of ca.
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4,000 Tn5-insertion mutants was constructed using colonies isolated from seven independent
rounds of electroporation. Ten colonies were randomly picked and the location of the
transposon insertion was investigated by Southern blot to rule out the presence of hot spots of
insertion in the genome and rates of single Tn5 insertions. To ensure the Tn5 insertion was
stably maintained in the absence of kanamycin, a randomly-selected mutant was transferred
for 16 and 58 days in kanamycin-free NBAF, then dilution plated on solidified NBAF medium
50

with and without KM , and the number of resulting colonies was compared.
High-throughput biofilm screening. Kanamycin-resistant colonies were transferred to
flat-bottom 96-well Costar cell culture cluster microtiter plates (Corning) containing 200 µl FW
30A/40F per well. The microtiter plates were incubated until visible turbidity was apparent (~35 days), and then transferred at least 3 times prior to inoculation of a biofilm assay. 10% (v/v) of
the culture in each well was then transferred to the well of a microtiter plated containing fresh
medium, and the plates were incubated at 30°C for 72 h. All steps were carried out in an
anaerobic glove bag (Coy), as described above. The plates were then removed from the glove
bag, the liquid culture with the planktonic cells discarded, and the biofilms were stained with
crystal violet, as described above. The plates were then allowed to dry in a fume hood
overnight and visually inspected to identify wells with less or more crystal violet staining than a
WT control well. The mutants having a distinct crystal violet staining were selected for a
quantitative secondary screening. Each mutant of interest was re-assayed in six replicates in
reference to the wild type. The crystal violet associated with the biofilm was dissolved in 33%
acetic acid for 20 min and the absorbance of the crystal violet solution was measured at 580 nm

100

on a SpectraMax M5 (Molecular Devices) or a Tecan Sunrise Plate Reader (Tecan, Inc.). Those
mutants with a reproducible biofilm phenotype (biofilm defective or enhanced) were
inoculated to a final OD600 of 0.04 in FW 30/40 medium in a 96-well plate and their growth at
o

30 C was monitored spectrophotometrically (OD600) every hour in a Tecan Sunrise plate
reader.
Identification of transposon-insertion sites. The transposon insertion site of mutants of
interest was investigated first via rescue cloning (26), and if unsuccessful, via direct genomic
sequencing (Britton Lab, personal communication), and/or arbitrary/touchdown PCR
techniques (6, 27, 40, 41, 44, 60). Genomic DNA (gDNA) was purified using the MasterPure DNA
Purification Kit (Epicentre Biotechnologies) according to the manufacturer’s specifications. All
sequencing reactions were performed at the Genomics Core of the Research Technology
Support Facility (Michigan State University).
Rescue cloning. 1 µg genomic gDNA was digested overnight with HindIII (New England
Biolabs) and self-ligated using T4 Ligase (Invitrogen) according to the manufacturer’s
+

specifications. The resulting plasmids were transformed into chemically-competent pir
50

Escherichia coli and plated on LB agar supplemented with KM . Colonies were picked and
grown in liquid LB with KM

50

before isolating the plasmid DNA using the ZR Plasmid Miniprep-

Classic Kit (Zymo Research). Sequencing of the Tn5 flanking regions was carried out using the
provided Tn5 reverse sequencing primer (KAN-2-RP-1). Sequences of primers used in this work
are listed in Table 4.1.

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TABLE 4.1: Primers
Primer Name
Sequence
KAN-2-FP-1
5'-ACCTACAACAAAGCTCTCATCAACC-3'
KAN-2-RP-1
5'-GCAATGTAACATCAGAGATTTTGAG-3'
ARB1
5'-GGCCACGCGTCGACTAGTACNNNNNNNNNNGATAT-3'
ARB2
5'-GGCCACGCGTCGACTAGTAC-3'
ARB6
5'-GGCCACGCGTCGACTAGTACNNNNNNNNNNACGCC-3'
APYBAF1
5'-CGGAATTCCGTGTTAAATATGGTATTGTGATNGAYKSNGGNTC-3'
HIB17
5'-CGGAATTCCGGATNGAYKSNGGNTC-3'
Tn5-F
5’-GCATTCACAGGGTGTCTCAA-3’
Tn5-R
5’-ATTCCGACTGGTCCAACATC-3’

Reference
(26, 27)
(26, 27)
(40, 41, 60)
(40, 41, 60)
(40, 41, 60)
(27)
(27)
This work
This work

Arbitrary PCR. An arbitrary PCR protocol was used, as described previously (41, 60).
Briefly, the first round of PCR uses a primer unique to the transposon (KAN-2-RP-1) and
arbitrary primers (ARB1 and ARB6) designed to hybridize randomly on the chromosome. The
second round of PCR coupled the KAN-2-RP-1 primer with a primer specific for the 5’ end of the
arbitrary primers (ARB2). All PCR reactions were performed on a Mastercycler epgradient S
(Eppendorf) using Go-Taq Green Mastermix (Promega) according to the manufacturer’s
specifications. The first round of PCR was carried out using the following conditions: 95°C for 2
min, followed by 30 cycles of 94°C for 30 s, 50°C for 30 s, and 74°C for 1 min, followed by a
final 5 min extension step at 74°C. The subsequent round of PCR amplification consisted of:
95°C for 2 min, followed by 5 cycles of 94°C for 30 s, 30°C for 30 s, and 74°C for 1 min, then 30
cycles of 94°C for 30 s, 30°C for 45 s, and 74°C for 1 min, and lastly followed by a final 5 min
extension step at 74°C. PCR products were run on a 1.2% agarose gel, stained with ethidium

102

bromide, visualized on an AlphaImager 2200 (Cell Biosciences) and bands were extracted using
the Zymoclean Gel DNA Recovery kit. Sequencing was carried out using the KAN-2-RP-1 primer.
For mutants that failed to produce a sequence using the above methods, alternative
arbitrary PCR methods were used, as described previously (27). The “touchdown” method of
arbitrary PCR involves an initial high annealing temperature (60°C), with a 0.5°C drop each
subsequent cycle, ending at 45°C. The Tn5-specific primers (KAN-2-FP-1 and KAN-2-RP-1) were
coupled with arbitrary primers APYBAF1 and HIB17. PCRs were performed and DNA bands
obtained as described above, except the following conditions were used: Touchdown protocol
1: 94°C for 8 min, followed by 30 cycles of 94°C for 1 min, 60°C-45°C for 1 min (each
temperature), and 72°C for 1 min. This was followed by 10 cycles of 94°C for 1 min, 45°C for 1
min, and 72°C for 1 min. Touchdown protocol 2 was carried out as follows: 95 °C for 5 min,
followed by 25 cycles of 95°C for 45 s, 60°C-45°C for 45 s (each temperature), and 72°C for 2
min. This was followed by 25 cycles of 95°C for 45 s, 50°C for 45 s, and 72°C for 2 min.
Mutant confirmation. Mutants of interest were colony purified, re-sequenced, and
subjected to Southern blotting to confirm the presence of a single Tn5 insertion. Genomic DNA
was extracted and digested with EcoRI Fast Digest Enzyme (Fermentas) according to the
manufacturer’s specifications. The digested DNA was run on an agarose gel and photographed,
before Southern blotting was performed as previously described (4). A probe of the Tn5 insert
was prepared by PCR amplifying the gene using primers Tn5-F and Tn5-R as well as DIG DNA
labeling mix (Roche) and Taq polymerase (Invitrogen) according to the manufacturer’s
103

recommendations with an annealing temperature of 55 °C. Detection was carried out using the
anti-digoxigenin-AP fab fragments (Roche) antibody and NBT/BCIP tablets (Roche).
SDS-PAGE and heme-stain of proteins in biofilm EPS matrix. EPS was extracted from
biofilms grown for 72 h on 6-well plates using the protocol described in previous published
works (8, 49), as well as in chapter 3 of this dissertation. Total protein content was quantified
using the Pierce Microplate BCA Protein Assay Kit (reducing reagent compatible, Thermo
Scientific) with BSA standards, according to the manufacturer’s specifications. Protein was
measured as an OD562 on a Tecan Sunrise Plate Reader.
For analyses of heme-containing proteins in the biofilm EPS matrix, 20 µg of protein
from each sample was boiled for 10 min in Lamaelli loading buffer (BioRad) and run at 250 V for
30 min on a 12% Mini-Protean TGX gel (BioRad). Heme-containing proteins were identified
using N,N,N’,N’-tetramethylbenzidine staining (11, 54). A duplicate gel was run in parallel and
stained for total protein using Coomassie Brilliant Blue R-250 (BioRad) according to the
manufacturer’s recommendations. The relative molecular weight of the proteins was estimated
using the Novex Sharp molecular weight markers (Invitrogen) as a standard.
U(VI) resting cell suspension biofilm assays. The ability of cells to remove U(VI) from
solution was assayed in resting biofilm cell suspensions using protocols adapted from those
described previously (11, 51), and described in Chapter 3 of this dissertation. Uninoculated
controls were also included to rule out any abiotic U absorption. Resting biofilm cell
suspensions were prepared from biofilms grown for 72 h in FW 30A/40F on sterilized 60 mm
glass petri dishes (Corning). The culture broth was carefully removed and the biofilms were

104

rinsed once with sterile, anaerobically-prepared wash buffer (51). This solution was replaced
with reaction buffer (51) supplemented with 20 mM sodium acetate and 1 mM uranyl acetate
(Electron Microscopy Sciences) prepared in 30mM bicarbonate buffer. The cell suspensions
were incubated for up to 24 h at 30°C under strictly anaerobic conditions. The U removal
activity was monitored periodically in 500 L samples. The samples were filtered (0.22 µm
Millex-GS filter, Millipore), acidified in 2% nitric acid (500 L), and stored at -20°C. The
concentration of U(VI) remaining in solution in the acidified samples was measured using a
Kinetic Phosphorescence Analyzer (KPA) (Chemchek).

RESULTS AND DISCUSSION
Development of an assay to characterize biofilm developmental stages. We
investigated biofilm formation in G. sulfurreducens using several commercially-available 96-well
plates. From all the microtiter plates tested, the tissue-culture-treated plate (Costar Cell Culture
Cluster) supported the formation of the most robust biofilms (Fig. 4.A.1). Other chemical
coatings such as Sigmacote (Fisherbrand plates coated), 0.01% poly-D-lysine hydrobromide, or
graphite were not as effective. Biofilm formation was, for example, prevented in the graphitecoated plates, whereas Sigmacote and poly-D-lysine-coated plates resulted in biofilms with
biomass ~80% of that measured in the uncoated control plates (Fig. 4.A.1). These differences in
binding could be attributed to the hydrophobic and ionic interactions between the bacteria and
the surfaces tested. The uncoated polystyrene and PVC 96-well plates (Fisherbrand, BD Falcon,
Nunc MicroWell) are hydrophobic and neutrally charged, and have been used in previous

105

biofilm assays (40, 41, 48). Graphite is also hydrophobic, whereas Sigmacote is a neutral and
hydrophobic film coating prepared from chlorinated organopolysiloxane in heptane. Thus,
hydrophobic surfaces are unlikely to favor the cell-surface interactions needed in the first
stages of biofilm formation. Poly-D-Lysine, on the other hand, is hydrophilic and positively
charged. It is commonly used to promote the attachment of cells or molecules with a
predominantly negative surface charge (Corning microplate selection guide, 2011; Corning
assay surfaces, 2007). However, biofilm formation was similar in tissue-culture treated plates
(Nunc MicroWell MaxiSorp, Corning Costar), which provide a hydrophilic and negativelycharged surface to promote the attachment of positively charged biomolecules (Corning
microplate selection guide, 2011; Corning assay surfaces, 2007). Thus, the charge of the surface
does not appear to affect cell attachment. The best performing plate, the Costar Cell Culture
Cluster, is a tissue-culture-treated plate that presumably provides a hydrophilic surface for cell
attachment; however, information about the chemical nature of the coating is not available. It
promoted biofilm formation in G. sulfurreducens and was selected for further studies.
Four solvents are commonly used to de-stain crystal violet biofilms (Table 4.A.1) (37),
and each was investigated for their suitability for our biofilm system. The maximum absorbance
reported for these solvents (590nm for DMSO, 580nm for all others) was used to quantify the
amount of crystal violet that was solubilized from G. sulfurreducens biofilms. All of the solvents
effectively solubilized most of the biofilm-associated dye, with ethanol and acetic acid
producing the highest absorbance values (Fig. 4.A.2). As ethanol evaporates faster and this can
concentrate the crystal violet solution, we chose acetic acid as a solvent to solubilize the crystal
violet associated to G. sulfurreducens biofilms.
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The first biofilm assays described for G. sulfurreducens used glass coverslips and FW
15A/40F medium (47), which we used above to screen for surfaces suitable for biofilm assays.
More recently, minimal media (DB) with excess electron donor (DB 30A/40F) was reported to
enhance biofilm attachment in 96-well plates (48). This is not surprising as previous studies
have noted the response of biofilms to nutritional cues (13, 41, 42, 61). Under the conditions of
our biofilm assay, FW 15A/40F promoted biofilm formation more rapidly than the DB 30A/40F
but the total biofilm biomass was similar in both (Fig 4.1A). However, providing an excess
electron donor to the FW medium (FW 30A/40F) promoted biofilm formation and resulted in
maximum biofilm growth at 72 h (Fig. 4.1A).

FIGURE 4.1: Comparison of the effect of growth medium type and electron donor (acetate)
concentrations on biofilm formation (A). CLSM images of wild-type biofilms grown in FW
30A/40F (B) (24 (top), 48 (middle), and 72 (bottom) h growth), FW 15A/40F (72 h growth) (C),
and DB 30A/40F (72 h growth) (D). Bar, 20 µM.

107

CLSM micrographs of the FW 30A/40F biofilms revealed three distinct biofilm structures at
24 h (monolayer), 48 h (microcolonies) and 72 h (mature biofilms) (Fig. 4.1B), consistent with
the developmental stages of initial attachment, microcolony formation, and maturation. The
three stages were also easily distinguished based on the crystal violet values shown in Fig.
4.1A). For these reasons, we selected FW 30A/40F as the growth medium for the purposes of
our screening.
Identification of Tn5-insertion mutants with biofilm defects. We used the Tn5
transposon to create a library of Tn5-insertion mutants. The transposon insertion was stable in
the absence of antibiotic (kanamycin) pressure, as approximately 2 weeks of serial transfers
50

yielded a 125% recovery on NBAF-KM

-

4

4

vs. NBAF-KM (5x10 CFU/ml vs. 4x10 CFU/ml), and
9

approximately two months of serial transfers yielded a 76% recovery (1x10 CFU/ml vs.
9

50

1.35x10 CFU/ml) on KM

media. Furthermore, Southern blot analyses of 10 mutant colonies

picked at random revealed a single transposon insertion site and at random locations (data not
shown), consistent with the random nature of the insertion and low likelihood of hot spots of
insertion. We then screened a Tn5 transposon-insertion library of 3,840 mutants to identify
mutants with both reduced and enhanced biofilm abilities. A primary visual screening identified
308 mutants. Of those, only 166 gave reproducible biofilm phenotypes in a quantitative
secondary screen. Out of the 166 mutants, 13 had enhanced biofilms and were not investigated
further. The remaining mutants have various degrees of biofilm defects and were grouped in
categories based on their crystal violet values: “no attachment” (“-“, interrupted at the
attachment stage of development, <5% of wild-type biofilms), “low” biofilm-formers (“+”,
108

corresponding to the microcolony stage, 5-40% of wild-type biofilms), and “medium” biofilmformers (“++”, interrupted at the maturation stage, 40-80% of wild-type biofilms) (crystal violet
de-staining values of representative mutants from each developmental stage are shown in Fig.
4.2).

FIGURE 4.2: Crystal violet staining of representative Tn5-insertion mutants. Biofilm crystal
violet staining values of in reference to the wild-type (left) and images (right) are shown. -, no
attachment; +, low biofilm; ++, medium biofilm; +++, wild-type.

Description of mutant groups. Table 4.A.2 provides an exhaustive list of the biofilmdeficient mutants whose insertion site could be identified, the interrupted gene locus, the
coordinates of the transposon insertion site, and growth phenotype. We identified genes from
numerous functional categories including electron transfer, cell envelope biogenesis, signal
transduction/regulation, metabolism, hypothetical/conserved hypothetical and unknown
function.

109

Cell envelope. Cell envelope components identified in this screening that have previously
been linked to biofilm formation in other bacteria include GSU3343 (SpoVR family protein, low
biofilm), which has been hypothesized to be responsible for synthesis of cell wall peptidoglycan
(5). Other examples include GSU1970 (polysaccharide biosynthesis protein, putative, low
biofilm), which is homologous to neuB in G. metallireducens and other organisms, as well as
GSU3460 (glycosyl transferase, group 2 family protein, medium biofilm). Both of these genes
has been previously implicated in the synthesis of capsule polysaccharides (2, 16, 55, 58). In
general, it appears that the expression of cell envelope biogenesis genes is required mainly for
the early stages of biofilm formation, consistent with their role in promoting productive cellsurface interactions.
Interestingly, many of the genes identified in the regulation, metabolism, and unknown
function categories were related to the biogenesis of the cell envelope, as well as electron
transfer. Examples of this include no attachment mutant GSU3385 (phophoenolpuruvate
carboxykinase, pckA), which is involved in the conversion of oxaloacetate to
phophoenolpyruvate and CO2 during the TCA cycle. Inactivation of this gene has been shown to
prevent capsule biosynthesis in Staphylococcus epidermidis (50). Other metabolic genes include
low biofilm-forming mutant GSU1942 (UDP-glucose/GDP-mannose dehydrogenase family
protein), which has been implicated in EPS and capsule biosynthesis (38, 56), as well as medium
biofilm-former GSU3321 (phosphoglucomutase/phosphomannomutase family protein) which is
responsible for the formation of alginate and therefore the production of LPS in Pseudomonas
aeruginosa. Expression of alginate has been correlated with the ability of cells to remain
attached to the substratum during the biofilm developmental process (18). Similarly, GntR-like
110

regulators (GSU1626, low biofilm-former) have been identified in an EPS gene cluster in
Streptomyces spp. (57). None of these mutants had defects in growth rates. Thus, they are
likely involved in the synthesis of a cell surface capsule or EPS matrix that is necessary for cell
attachment.
Electron transfer. We identified genes predicted to regulate, directly or indirectly, the
expression and functioning of electron transport components.
These genes included those involved in the biogenesis and regulation of the conductive pili,
c-type cytochromes, and hydrogenases. Mutants carrying Tn5 insertions in c-type cytochromes
were identified in the low and medium biofilm categories, suggesting that they are required for
the development of multilayered biofilms and their maturation. Among the genes we identified
several were unable to form microcolonies (GSU2912, GSU0274, and GSU3332, all cytochrome
c family proteins), or mature biofilms (GSU0222, cytochrome c oxidase, subunit II). Information
about these cytochromes is scarce. Mutants in GSU2912 are defective in the reduction of
soluble Fe(III) citrate, but not the insoluble Fe(III) oxides (48). Mutants in GSU3332, a predicted
outer membrane c-cytochrome, have defects in both uranium and Fe(III) oxide reduction (51),
consistent with their role in extracellular electron transfer. No information is available for
GSU0274 or for the cytochrome c oxidase of G. sulfurreducens. However, cytochrome c oxidase
has previously been shown to be required for the later stages of biofilm formation in
Pseudomonas aeruginosa, potentially because it is required for growth in anoxic environment,
and oxygen gradients are common to the later stages of biofilm formation, such as the
development of microcolonies (52).

111

Among the genes known to have an indirect effect, we identified one encoding a
CRISPR-associated protein, cas2 (GSU0058, low biofilm), which has been hypothesized to inhibit
c-type cytochromes in Pelobacter carbinolicus, a close relative of G. sulfurreducens (1). Another
gene of potential interest in electron transfer is the fibronectin type III domain protein
(GSU1945, medium biofilm). Although annotated as a protein of “unknown function”,
fibronectin domains are often linker domains that promote the binding and catalyses of
substrates. Some cellulases, for example, have fibronectin domains to connect the catalytic and
binding domains and promote the binding of the enzyme to cellulose fibers and its hydrolysis,
presumably because it directs the cellulose into the active site and maintains the optimal
interaction and orientation between it and the catalytic domain of the glycosyl hydrolase (24,
59). A similar function has been proposed for the fibronectin domain in the OmpB protein, a
multicopper protein of G. sulfurreducens that promotes the association of the cell with
insoluble Fe(III) oxides to facilitate electron transfer (36).
The mutants with the transposon inserted in pili biogenesis genes ranged from no
attachment (GSU1493, pilC), low (GSU1492, pilR), and medium (GSU1495, pilT-4) biofilmformers. This suggests that pili expression and function is required throughout the
development of the biofilms, consistent with their role as structural components of the biofilm
matrix and electronic conduits across the biofilms (46, 47). Similar genes homologues have
been linked to the development of mature biofilms in other organisms (40, 41, 43-45, 47). In G.
sulfurreducens, mutations in the pilR gene reduce the transcription of the pilA gene, which
encodes the pilin subunit, as well as several c-cytochromes (23). As a result, a pilR mutant has
defects in pili expression and cell-aggregation (23), which can lead to defects in biofilm
112

formation. Interestingly, although a pilT-4 mutant was biofilm-deficient in our screening, it was
previously reported to have enhanced biofilm capabilities (48). The biofilm screening used in
this early report used DB 30A/40F medium, which we showed does not promote the growth of
multilayered biofilms (Fig. 4.1A). Thus, the discrepancies in the reported phenotypes may have
resulted from the different growth media used in both assays. The pilT-4 gene is one of 4 pilT
homologues identified in the genome of G. sulfurreducens. In other bacteria, PilT is an ATPase
that powers pilus retraction and depolymerization (35, 62). As a result, mutations in pilT often
lead to hyperpiliated strains and enhanced biofilm abilities (35). The biofilms formed by
hyperpiliated strains often have tall mushroom-like pillars (9), which can easily break during the
biofilm assay, thus leading to a biofilm-defective phenotype. Alternatively, pilT-4 may not
encode for a functional PilT ATPase in G. sulfurreducens or in the biofilms. We also identified a
“no attachment” mutant with a transposon insertion in the pilC gene. PilC is an integral
membrane protein associated with the pilus apparatus, and has been proposed to be involved
in secretion, assembly, and/or stabilization of the pilus fiber (62). However, its role in pilus
biogenesis has not been conclusively demonstrated. The finding that a pilC mutant cannot
attach to the surface to initiate biofilm suggests that the pili of G. sulfurreducens could also be
involved in the early stages of biofilm formation, when the cell attaches to the surface as a
monolayer. However, the phenotype of the pilC mutant was variable, with some independent
assays showing no attachment and others (Fig. 4.3A) showing medium biofilm formation. These
differences could reflect differences in the growth rates or surface properties of the mutant
cells in independent experiments, which are not uncommon in pili-deficient mutants in G.
sulfurreducens (11). Thus, the results do not conclusive link pili expression to attachment.
113

Role of pili in biofilm development and uranium immobilization. We gained further
insights into the role of pili in biofilm development by investigating the phenotypes of the
three strains carrying mutations in pili genes: pilC::Tn5 (GSU1493, integral membrane protein,
54

pili biogenesis), pilR::Tn5 (GSU1492, σ -dependent response regulator), and pilT-4::Tn5
(GSU1495, twitching motility protein PilT). The mutants were purified, their insertion site reconfirmed by sequencing, and a Southern blot was performed to demonstrate that the
observed phenotype was the result of a single insertion event (Fig. 4.A.3). A new biofilm assay
confirmed the biofilm defects in the pilR and pilT-4 mutants; however, the pilC::Tn5 mutant
attached to the surface but could not develop microcolonies (Fig. 4.3A). The uranium removal
2

activities of the biofilms correlated well (R = 0.90) with the amount of biofilm biomass (Fig.
4.3B), suggesting that pili expression during biofilm maturation is necessary to maximize the
removal potential of the biofilms.
As mutations leading to changes in pili expression can also result in defects in the
expression of outer membrane c-cytochromes (11) and the cytochromes of the biofilm matrix
(Chapter 3), we extracted the EPS matrix from the mutant biofilms and separated the proteins
associated with the EPS matrix electrophoretically. The protein in the gel was then stained to
identify the heme-containing proteins. The heme content and profile of the pilC::Tn5 mutant
was similar to the WT (Fig. 4.3C). This is significant because all the mutations in pili biogenesis
genes investigated thus far (11, 23) also lead to defects in c-cytochrome expression. Thus,
defects in metal reduction in these mutants are difficult to interpret because of the pleiotropic
nature of the mutations. Thus, this gene is a good molecular marker to link the activity of the

114

biofilms conductive pili to its reductive capabilities. By contrast, both pilR::Tn5 and pilT-4::Tn5
had defects in heme-containing proteins, including the OmcZS c-cytochrome (~30 KDa) (Fig.
4.3C), which we previously correlated with the uranium reductive activities of G.
sulfurreducens biofilms (chapter 3). The defects observed in the pilR::Tn5 are consistent with
previous reports that demonstrated the differential regulation of a number of c-type
-

cytochromes in a PilR mutant (23). The pilT-4::Tn5 mutant biofilms also had a reduced heme
content in the EPS matrix (Fig. 4.3C). As pili retraction leads to hyperpiliated strains (35), it is
possible that overproducing pili affects c-cytochrome secretion, as we previously show for the
hyperpiliated pRG5::pilA strain (11), which expresses the pilA gene from a medium-copy
-

plasmid (45). It is interesting to note that planktonic cells of a PilT-4 mutant do not have any
defect in the expression of outer membrane c-cytochromes (Bryan Schindler, unpublished
data).

115

FIGURE 4.3: Phenotypic characterization of selected mutants. Biofilm assay (A), correlation between U removal and biomass (B),
and heme content of EPS (C) are shown. Lane 1, WT; lane 2, pilC::Tn5; lane 3, pilR::Tn5; lane 4, pilT4::Tn5.

116

Conclusions. The results demonstrate that biofilm formation in G. sulfurreducens is a
developmental process requiring the expression of specific genes at each stage. Genes
encoding proteins for capsule and EPS synthesis were required in the early stages of
attachment to form monolayered biofilms, consistent with the role of specific components of
the cell’s exterior at promoting cell-surface interactions. The conductive pili of G.
sulfurreducens were required for the formation of microcolonies and for biofilm maturation,
thus providing additional evidence for the requirement of pili expression as structural support
and electronic connections in multilayered biofilms (46, 47). Interestingly, c-cytochromes were
also required for the formation of microcolonies and biofilm maturation. As the biofilms
increase in height, c-cytochromes in the biofilm EPS matrix promote long-range electron
transfer across the biofilms, thereby supporting the growth of the biofilm cells (28). Taken
together, the genetic analyses of biofilm development in G. sulfurreducens revealed
conservation of components required for biofilm formation in other organisms but also unique
components required for the electroactivity of the biofilms.

117

APPENDIX

118

TABLE 4.A.1: Plates, coatings, growth media, and solvents assessed for their use in the biofilm
assay.
96-Well Plate
Manufacturer & Catalog Number
Nunc MicroWell Plate
Thermo Scientific Cat. No. 260836
Nunc MicroWell MaxiSorp Plate
Thermo Scientific Cat. No. 456537
Costar Cell Culture Cluster Plate
Corning Cat. No. 3595
Fisherbrand Clear Polystyrene Plate
Fisher Scientific Cat. No. 15-565-500
BD Falcon PVC Flexible U Bottom Plate
BD Biosciences Cat. No. 353911
Coating
Sigmacote
0.01% Poly-D-Lysine Hydrobromide
Graphite (Ladd Carbon Evaporator)

Manufacturer & Catalog Number
Sigma-Aldrich Cat. No. SL2
Sigma-Aldrich Cat. No. P7886
Ladd Research

Growth Medium

Reference

FW 15A/40F
FW 30A/40F
DB 30A/40F

(29), with modifications described in (11)
This work
(33, 48, 53)

Solvent

Absorption Wavelength

100% DMSO
95% Ethanol
33% Acetic Acid
80% Ethanol: 20% Acetone

590nm
580nm
580nm
580nm

119

FIGURE 4.A.1: A selection of 96-well plates investigated for their ability to maximize G. sulfurreducens 72 h biofilm growth at
30°C.

120

FIGURE 4.A.2: Selection of a solvent for de-staining of crystal violet biofilms. A comparison of
the ability of four solvents to dissolve crystal violet-stained biofilms. Shown here is the optical
density read at the optimal absorbance for each solvent (DMSO, 590nm; all others, 580nm).

121

TABLE 4.A.2: Transposon-insertion mutants displaying deficient or enhanced biofilm phenotypes
GSU

a

Growth
Rate
+
+
+
-

Biofilm
-

+
+

+
+

3661881-3661882
3785649-3785650
3043161-3043162

+
+
+

+
+
+

2011467-2011468
1330619-1330620
1573183-1573184
3695163-3695164
262719-262720
3671647-3671648
1781558-1781559
3420238-3420239

+
+
+
+
+
+
+
+

+
+
+
+
+
+
+
+

310083-310084
505038-505039
1641425-1641426

+
+
+

+
+
+

c

locus
2282
1945
3385
1493

CBS Domain Protein
Fibronectin type III domain protein
Phosphoenolpyruvate Carboxykinase (pckA)
Type IV Pilus Biogenesis Protein PilC

24 D5
21 B1

2912
0274

Cytochrome C Family Protein
Cytochrome C Family Protein

7 G8
25 E1
21 A11

3332
3439
2762

35 E2
19 C10
23 C2
17 C6
19 H10
20 G1
21 B2
21 D4

1843
1227
1434
3363
0253
3343
1626
3118

23 D1
27 E10
35 B2

0281
0474
1495

e

Annotation

b

Mutant
21 A2
22 A10
29 B10
31 C5

Cytochrome C Family Protein
NADH Dehydrogenase I, G Subunit
Glycerol Kinase (glpK)
Metallo-beta-lactamase family protein (similar to RNA
metabolizing)
ABC Transporter, ATP-Binding Protein
Peptide ABC transporter permease protein
Sigma-54-Dependent Transcriptional Regulator, Fis Family
Sensory Box Histidine Kinase
SpoVR-Like Family Protein
Transcriptional Regulator, GntR Family
DNA-Binding Response Regulator
Sensor histidine kinase, authentic frameshift (near cadherin
gene)
Sensory Box/GGDEF
Sigma-54 dependent DNA-binding response regulator
122

Coordinate
2498835-2498836
2132741-2132742
3723531-3723532
1638348-1638349
320949732099498
283025-283026

d

TABLE 4.A.2 (cont’d)
25 B3
0058 CRISPR-associated protein, cas2 (298bp after –sRNA region)
22 H2
0002 RecF Protein
21 A4
2387 B12-Binding Domain
21 E9
3157 Hydrolase, alpha/beta fold family
e

23 H12
25 C2
19 H11
21 A8
35 A3
35 B6
35 F6
35 G6
35 B7
35 C7
37 D2

0978
0238
1261
1700
0474
0474
1122
0992
3199
0174
1074

37 D9
37 F2

2403
1949

39 F11
39 H1
3 H11
7 A11
7 C12
1D9
1 C8

1942
1970
3295
0474
1485
1183
2502
3459/

3 A1

3460

e

e

74899-74900
3030-3031
2616527-2616528
3464948-3464949

+
+
+
+

+
+
+
+

Hypothetical Protein (in Phage Operon)
Radical SAM Domain Protein
ATP transporter, ATP binding protein
NAD-dependent malic enzyme (maeB)
Sensory Box/GGDEF
Sensory Box/GGDEF
HD Domain Protein
Hypothetical protein
Chemotaxis protein CheA (CheA-3)
Acetyl-CoA Hydrolase/Transferase Family Protein
DNA-Binding Regulatory Protein, YebC/PmpR Family

1055246-1055247
246566-246567
1373553-137354
1862353-1862354
505038-505039
505038-505039
1207423-1207424
1073609-1073610
3507763-3507764
189971-189972
1163749-1163750

+
+
+
+
+
+
+
+
+
+
+

+
+
+
+
+
+
+
+
+
+
+

Hypothetical Protein
Hypothetical Protein
UDP-Glucose/UDP-Mannose Dehydrogenase Family
Protein
Polysaccharide Biosynthesis Protein, Putative
Conserved Hypothetical Protein
Sensory Box/GGDEF
Ribonuclease R, putative
O-acetyl-L-homoserine sulfhydrylase
Spermine/Spermidine Synthase Family Protein

2638633-2638633
2136757-2136758

+
+

+
+

2127220-2127221
2161395-2161396
3614122-3614123
505038-505039
1628979-1628980
1290628-1290629
2756295-2756296

+
+
++
+
+
+
+

+
+
+
+
+
++
++

3804868-3804869

+

++

Hypothetical Protein & Glycosyl transferase, group 2 family
protein (between)
123

TABLE 4.A.2 (cont’d)
17 H7
1432 TRP Domain Protein
21 A5
0377 Glycine Cleavage System, P Protein, Subunit 1
Glycine Cleavage System Transcriptional Repressor,
26 H8
3457 Putative
21 A9
0775 Sensor Histidine Kinase
21 B12
1337 Lipoprotein, Putative
Phosphoglucomutase/Phosphomannomutase Family
21 C5
3321 Protein
21 F11
3399 Efflux Transporter, RND family, MFP Subunit
21 F12
0222 Cytochrome c oxidase, subunit II
2319/2 Conserved Hypothetical (Upstream of CFA Operon) (245bp
e

22 H6
23 E1
23 E2
23 F1
24 A3
24 A7
24 B4

320
1672
0180
0968
3318
1185
3418

24 E2
24 F8
24 G2

3326
2697
1945

25 A8
25 D11
25 F6
25 H10
26 B12

0098
0238
2671
1492
0306

e

1569923-1569924
411286-411287

+
+

++
++

3804144-3804144
832499-832500
1464937-1464938

+
+
+

++
++
++

3647205-3647206
3739015-3739016
230554-230555

+
+
+

++
++
++

upstream), Hypothetical (9bp upstream)
Glycerate dehydrogenase (hprA)
Conserved Hypothetical Protein (near metabolic genes)
Hypothetical protein
Conserved Hypothetical Protein
Conserved Hypothetical Protein
Sigma 54-Dependent DNA-Binding Response Regulator
Conserved Hypothetical (Downstream of Exinuclease ABC
(uvrA))
Multidrug Resistance Protein
Fibronectin type III domain protein

2540037-2540038
1833040-1833041
194573-194574
1042230-1042231
3644746-3644747
1293096-1293097
3761837-3761838

+
+
+
+
+
+
+

++
++
++
++
++
++
++

3655240-3655241
2978002-2978003
2132741-2132742

+
+
+

++
++
++

MglB Protein (29bp upstream)
Radical SAM Domain Protein
Hypothetical Protein
Twitching Motility Protein PilT (PilT-4)
Hydrogenase Maturation Protein (HypF)

113579-113580
246566-246567
2946824-2946824
1637002-1637003
339834-339833

+
+
+
+
+

++
++
++
++
++

124

TABLE 4.A.2 (cont’d)
26 H1
0302 Hypothetical Protein
28 G12
3158 Cysteine Synthase B (cysM)
3459/3 Hypothetical Protein & Glycosyl transferase, group 2 family
e

35 C3

460

35 E8
35 F8
35 H10
35 G11

1733
0728
1358
1009

38 A8
38 C5

0323
1527
2330/2
332
3386
1007
0244
0924
3459/3
460
2330/2
332
0058

28 B8
28 B11
28 H10
29 H10
31 B9
32 D9
32 F12
1D1
a

e

331689-331690
3465380-3465381

+
+

++
++

protein (between)
Branched-chain amino acid ABC transporter permease
protein (livH)
Polyphosphate:AMP Phosphotransferase (pap)
Conserved Hypothetical Protein
GTP-Binding Protein
General Secretion Pathway Protein J, Putative (47bp
upstream)
Conserved Hypothetical Protein

3804868-3804869

+

++

1898384-1898385
775630-775631
1484097-1484098
1090260-1090261

+
+
+
+

++
++
++
++

354290-354291
1674377-1674378

+
+

++
++

Conserved Hypothetical and Hypothetical (CFA Operon)
Lipoprotein, putative
GAF Domain/HD Domain Protein
Radical SAM Domain Protein
ABC Transporter, Permease Protein, Putative
Hypothetical Protein & Glycosyl transferase, group 2 family
protein (between)

2550521-2550522
3723753-3723754
1087571-1087572
251774-251775
993660-993661

+
+
+
+
+

++++
++++
++++
++++
++++

3804868-3804869

+

++++

Conserved Hypothetical and Hypothetical (CFA Operon)
CRISPR-associated protein, cas2 (298bp after –sRNA region)

2550521-2550522
74899-74900

+
+

++++
++++

b

c

GSU locus containing the transposon insertion, Annotation assigned by JCVI, Genomic coordinate for the transposon insertion

d

e

Biofilm phenotype: no attachment (-), low (+), and medium (++) in reference to WT (+++), Insertion just upstream of designated
open reading frame

125

FIGURE 4.A.3: Southern blot to confirm a single Tn5 insertion event. A positive control of the
Tn5 probe (+) and negative control (wild-type genomic DNA) (-) are shown in addition to the
pilC::Tn5, pilR::Tn5, pilT-4::Tn5 insertion mutants (2-4 clones of each).

126

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CHAPTER 5

IDENTIFICATION AND CHARACTERIZATION OF THE RPOE SIGMA FACTOR OF
GEOBACTER SULFURREDUCENS

The following former members of Dr. Gemma Reguera (Michigan State University) and Dr.
Derek Lovley’s (University of Massachusetts, Amherst) laboratories contributed to the work
presented in the chapter:

Blair Bullard (Michigan State University): Biofilm assays
Dr. Evengenya Shelobolina (University of Massachusetts, Amherst): resting cell suspensions
Dr. Gemma Reguera (University of Massachusetts, Amherst/Michigan State University): mutant
generation, bioinformatic analyses, biofilm assays, and microarray analyses

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ABSTRACT
Metal and electrode reduction in Geobacter bacteria is intimately connected to the
integrity and functionality of the cell envelope, yet little is known about its regulation. Here we
report on the genetic identification and functional characterization of the RpoE sigma factor of
Geobacter sulfurreducens (RpoEGsu). As reported for other RpoE sigma factors such as in
Escherichia coli, RpoEGsu regulated growth transitions in response to nutritional shifts and
during biofilm formation, and was also involved in the oxidative stress response. All of these
functions ultimately depended on the role of RpoE in the regulation of cell envelope integrity
and homeostasis. Despite these conserved roles, RpoEGsu was not required for growth at high
temperatures or low pHs, but for growth at lower temperatures and higher pHs, thus reflecting
the adaptation of G. sulfurreducens extracytoplasmic functions to stressors commonly found in
environments inhabited by Geobacter bacteria. The phenotypes were also consistent with
transcriptomic profiling, which identified genes in the RpoE regulon required for conserved
mechanisms of RpoE response to cell envelope stress as well as specialized responses required
to regulate Geobacter extracellular electron transfer pathways. This work highlights the
functional specialization that RpoE has undergone to control the adaptive responses that
enable Geobacter bacteria to survive in the subsurface and that are relevant for applications in
bioremediation and current generation in microbial fuel cells.

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INTRODUCTION
Microorganisms in the family Geobacteraceae conserve energy to support growth from the
reduction of Fe(III) and Mn(IV) (44) and are also recognized as important agents in the
bioremediation of organic (2, 43, 65, 66, 70) or metal (3, 46, 60) contaminants, and as primary
contributors to current production in microbial fuel cells harvesting electricity from the
environment (4, 23, 42, 71). Key to their physiology and applications is the control of electron
transfer across the cell envelope. The genome sequence of some representative species of the
Geobacteraceae available in pure culture reveals a great number of genes potentially encoding
cell envelope proteins involved in electron transfer processes as well as a complex regulatory
circuitry that could be responsible for signal integration and response to environmental cues
(48). Studies in the model dissimilatory Fe(III) reducer Geobacter sulfurreducens have shown
that the expression of most genes known to be of relevance to electron transfer may be tightly
regulated (15, 24, 25, 28, 31-33, 35, 49, 57, 58, 73). Some of these studies (28, 34, 57, 58, 73,
76-78) have identified a great number of genes and operons encoding proteins relevant to
electron transfer and environmental survival that may be under transcriptional control by sigma
factors. The expression and/or availability of sigma factors is regulated by environmental
signals. As a result, sigma factors modulate the cell’s transcriptional machinery and allow it to
reprogram gene expression by ‘turning on’ those genes necessary to respond to a particular
environmental signal and ‘turning off’ those that are no longer needed (27).
The genome of G. sulfurreducens contains genes encoding homologues of the sigma
70

S

factors RpoD, RpoS, RpoH, RpoE and FliA of the  -family (48). The RpoS ( ) of G.
sulfurreducens controls the expression of genes required for survival in stationary phase,
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oxygen tolerance, heat shock and alkaline stress responses, and electron transfer pathways to
H

32

Fe(III) (57, 58), whereas RpoH ( ,  ) controls the heat shock response (73) and FliA is
predicted to function in regulating the expression of flagellar and chemotaxis-related genes (37,
72). To date, no functional characterization is available for RpoE in G. sulfurreducens.
RpoE is a member of the extracytoplasmic function (ECF) subfamily, which encompasses
70

those 

factors that respond to extracytoplasmic stimuli such as the accumulation of

misfolded proteins in the periplasmic space (RpoE) or the extracellular availability of ferric
citrate (FecI) (16, 22, 52, 62). The RpoE homologue of G. sulfurreducens is the only ECF sigma
factor identified in its genome (48). Although annotated as a putative RpoE sigma factor,
sequence analyses of the encoding gene failed to shed light on its function, and putative roles
as an RpoE (57) or a FecI (76) sigma factor have been proposed. Bioinformatics analyses using E.
coli’s known transcription factors and transcription regulatory sites also failed to identify
conserved RpoE-like regulatory elements in the genome of G. sulfurreducens (76). As a result,
no functional confirmation is yet available for the putative RpoE sigma factor in this organism.
As this is the only ECF sigma factor homologue identified in the genome of G. sulfurreducens,
we hypothesized that the identified gene represents a functional RpoE sigma factor and major
regulator of cell envelope functions such as metal reduction and survival of this organism in the
subsurface. To further elucidate the role of RpoE, we investigated the genetic organization and
operon structure of the rpoE gene. We also studied the phenotype of an rpoE deletion mutant
under a variety of growth conditions and environmental stresses. The results demonstrate that
RpoE plays a role in the maintenance of cell envelope integrity and physiological responses of

137

Geobacter spp. that are relevant to metal reduction and applications in bioremediation and
microbial fuel cells.

MATERIALS AND METHODS
Bacterial strains and culture conditions. Geobacter sulfurreducens strain ATCC 51573
-

(6) was obtained from the American Type Culture Collection (ATCC). The RpoE-deficient (RpoE )
mutant used in this study was generated by complete deletion of the rpoE gene (GSU0721) and
insertion of a kanamycin resistance cassette in the same orientation of rpoE transcription
-

(Gemma Reguera, personal communication). The wild-type (WT) strain and RpoE mutant were
routinely cultured in NB (10) or fresh water (FW) (9, 45) medium containing 15 mM acetate and
40 mM fumarate (NBAF or FWAF). NBAF medium was supplemented with 1 mM cysteine-HCl
and 0.1% yeast extract and, when indicated, 20mM MOPS buffer. FWAF was supplemented
with 0.25 mM cysteine-HCl and 0.05% yeast extract. Cultures were incubated at 30°C unless
otherwise indicated. Growth curves were generated from mid-log phase cultures grown in
NBAF medium and inoculated to a final OD600 of 0.04 in NBAF medium. Growth was measured
spectrophotometrically by periodically monitoring the optical density (OD600) of the cultures.
Complementation. A wild-type copy of the rpoE gene was PCR-amplified using primers
RpoE-Comp-pRG5-F (BglII): GGAAGATCTAAGGAGAAGTGACGCTGGAT and RpoE-Comp-pRG5-R
(HindIII): CCCAAGCTTTCACGCTCGTCATGTTTCAT (restriction sites are underlined) and Phusion
high-fidelity DNA polymerase (New England Biolabs) according to the manufacturer’s
138

specification. PCRs were carried out on a Mastercycler Personal (Eppendorf) using the following
conditions: 98°C for 30 s, followed by 30 cycles of 98°C for 10 s, 55°C for 30 s, and 72°C for 30
s, followed by a final 10 min extension step at 72°C. The resulting PCR products were digested
with BglII and HindIII (New England Biolabs), and a medium-copy plasmid, pRG5 (32), was
digested with BamHI and HindIII (New England Biolabs) according to the manufacturer’s
recommendations. The digested products were ligated with T4 DNA ligase (Invitrogen)
-

according to the manufacturer’s recommendations. RpoE electrocompetent cells were
prepared as described previously (10), and ~1 µg of the pRG5::rpoE plasmid was electroporated
using a Eppendorf 2510 Electroporator operated at 1470 volts. Cultures were recovered
overnight and then plated on NBAF medium supplemented with 75 µg/ml spectinomycin to
select for the pRG5 plasmid. Controls included non-electroporated competent cells, cells that
were subjected to electroporation without the introduction of foreign DNA, and cells
electroporated with the pRG5 plasmid only (no insert).
Transcriptional analysis of the rpoE gene and the gene region by RT-PCR. Reverse
Transcriptase-Polymerase Chain Reaction (RT-PCR) was used to quantify the transcription of the
rpoE gene and the downstream genes (GSU0722 and GSU0723). For these experiments, cells of
-

the wild-type and RpoE mutant were grown in NBAF at 25°C, 30°C, and 35°C. RNA was
extracted using TRIzol reagent (Invitrogen), treated with RQ1 RNase-free DNase (Promega), and
reverse transcription was performed using Superscript III Reverse Transciptase (Invitrogen), all
according to the manufacturer’s recommendations. Reverse transcription and PCR
amplification of rpoE was performed using primers RpoE-qPCR-771279F (5’139

GTGACGCTGGATACAACAGACGAA-3’) and RpoE-qPCR-771430R
(5’GCGCATAGTTTCTGACAAAGCCGA-3’). The co-transcription of rpoE with the downstream
genes (GSU0722 and GSU0723) was studied using primer combinations RpoE-qPCR-771279 with
GSU0722-R (5’ TCATTGGCTGCAATCAACTC 3’) and GSU0723-R (5’ CCGGAATCATCAGTTGACCT
3’), respectively. A constitutively-expressed control gene, recA, was assayed using primers
recA660f and recA737r (25). Using the cDNA generated from reverse transcription, a PCR
reaction was performed on a Mastercycler epgradient S (Eppendorf) using the following
conditions: 95°C for 5 min, followed by 30 cycles of 94°C for 30 s, 55°C for 45 s, and 72°C for 1
min, followed by a final 10 min extension step at 72°C. The resulting products were run on a
1.2% agarose gel, stained with ethidium bromide, and visualized using an AlphaImager TM2200
(Cell Biosciences).
-

Stress tolerance assays. The aerotolerance of the wild-type and RpoE mutant strains
was evaluated in mid-exponential phase cultures grown in FWAF medium. 2 mL aliquots were
removed from the cultures and placed in sterile cuvettes capped with sterile aluminum foil and
incubated at 30°C. The OD600 of the cultures was monitored periodically and the aerotolerance
of the strains was inferred from the decreases in the cultures’ absorbance over time.
The effect of pH was assayed using an adaptation of a previously described method (57).
Briefly, cultures were grown to mid-exponential phase in MOPS-buffered NBAF medium (pH 7),
and then transferred (final OD600 of 0.04) to fresh medium buffered at pH 6 or 9. After 48 h (pH
6) and 24 h (pH 9) of exposure, the cultures were transferred (10% (v/v) inoculum) into

140

anaerobic NBAF medium (pH 7) and growth (OD600) was monitored periodically. The length of
the lag phase of the cultures was used to estimate the survival rates of the strains after
exposure to suboptimal pH conditions and, therefore, their pH tolerance
Biofilm assays. Cells were grown to mid-late exponential phase at 30°C in NBAF
medium and transferred three times prior to assaying for biofilm formation. Biofilm assays
2

were performed using glass coverslips (18x18 mm ), as previously described (64), except that
FWAF medium supplemented with 0.25 mM cysteine and 0.05% yeast extract was used as the
growth medium. Planktonic growth was monitored periodically by measuring the OD 600 of the
culture. Biofilm growth was quantified every 24 h by staining the biofilm biomass on the
coverslips with 0.1% crystal violet (59). The biofilm-associated dye was solubilized in 33% acetic
acid and quantified by measuring the OD580 of the dye-acid solution. The biofilms were also
washed in anaerobic PBS buffer and stained with the BacLight  viability kit (Invitrogen) prior
to examination by Confocal Laser Scanning Microscopy (CLSM) using a Zeiss 510 Meta ConfoCor
3 LSM equipped with a 40X objective.
Transmission Electron Microscopy (TEM). Early-stationary phase cultures grown in
NBAF medium were mixed with an equal volume of reaction buffer (67). The cell suspension
was adsorbed onto 300-mesh carbon-coated copper grids (Ted Pella), fixed with 1%
glutaraldehyde for 5 min, washed with ddH2O for 3 x 2 min, and stained with 1% uranyl acetate
for 30 s. Cells were imaged on a JEOL100CX electron microscope (Japan Electron Optic
Laboratory) operated at a 100 kV accelerating voltage.
141

Fe(III) reduction assays. The ability of the wild-type and the RpoE mutant strains to
reduce Fe(III) was measured in resting cell suspensions prepared as previously described (67).
Briefly, G. sulfurreducens strains were cultured in FW medium amended with 10 mM acetate
and 20 mM fumarate. Late logarithmic phase cultures were harvested by centrifugation and
washed twice in wash buffer, resuspended in reaction buffer, and Fe(III)-reducing activities
were quantified, as described before (67). Cells were added to the reaction buffer to give a final
OD of ca. 0.06-0.08. The ability of cells to reduce Fe(III) was determined using either poorly
crystalline ferric hydroxide (Fe(III) oxide) (20 mM) or ferric citrate (Fe(III) citrate) (20 mM) as the
electron acceptor and 5 mM acetate as the electron donor. Fe(II) concentrations were
determined hourly using a ferrozine assay as previously described (69). All Fe(III) reduction
assays were performed by Evgenya Shelobolina (University of Massachusetts, Amherst).

RESULTS AND DICUSSION
Genetic organization of the rpoE region. ECF sigma factors are often co-transcribed
with a transmembrane antisigma factor and an accessory protein, such as RseA and RseB in E.
coli, respectively (13) . The RseA antisigma factor of E. coli, for example, is an inner membrane
E

protein with a periplasmic sensory domain and a cytoplasmic anti- domain that sequesters
the RpoE protein, thus, controlling its cytoplasmic availability and binding to the RNA
polymerase (7, 13, 51). The inhibitory function of RseA is enhanced by RseB, a periplasmic
protein that binds to RseA’s periplasmic domain and protects it from proteolysis (1, 13, 21, 51).
Although a search for RseA and RseB homologues in the genome of G. sulfurreducens using E.
coli’s amino acid sequences retrieved no matches (Gemma Reguera, personal communication),
142

bioinformatics analyses (76) predicted that the gene encoding RpoE in G. sulfurreducens (herein
termed RpoEGsu) is part of a predicted operon with genes encoding a hypothetical protein
(GSU0722) and a conserved hypothetical protein (GSU0723) (Fig. 5.1A).

FIGURE 5.1: Genetic arrangement of the rpoE region (A) and transcriptional analysis by reverse
transcriptase (RT) PCR of the predicted operon (B). Controls with (+RT) or without (-RT) reverse
transcriptase are shown for the rpoE operon transcripts, as well as controls with WT genomic
DNA (gDNA). 1, GSU0721 only; 2, GSU0721+GSU0722; 3, GSU0721+GSU0722+GSU0723.

The hypothetical protein encoded by GSU0722 contained one internal helix between
amino acids 55 and 74 that could function as a transmembrane domain and had no predicted
signal peptide. This is consistent with the cytoplasmic membrane localization expected for a
putative antisigma factor. The conserved hypothetical protein encoded by GSU0723 contained
a signal peptide with a putative cleavage site between amino acids 35 and 36 (AWG-DG) and
143

had no transmembrane domains, suggestive of a periplasmic localization, analogous to RseB of
E. coli. However, transcriptional analyses of the rpoE gene region demonstrated that rpoE is not
co-transcribed with the downstream genes under the conditions tested. Rather, rpoE is
transcribed independently of the two downstream genes (Fig. 5.1B). These results can be
interpreted as RpoEGsu being regulated by mechanisms that differ substantially from the
canonical regulation of RpoE in E. coli and other bacteria, similarly to what has been proposed
for several mycobacterial species (74). Interestingly, the genetic arrangement of the rpoE gene
homologue in G. sulfurreducens is conserved in other Geobacter spp. such as Gebacter
metallireducens and Geobacter uraniireducens, both containing genes downstream of the rpoE
homologue encoding a hypothetical protein (45% identity out of 121 amino acids and 38%
identity out of 118 amino acids, respectively) and a conserved hypothetical protein (55%
identity out of 134 amino acids and 42% identity out of 146 amino acids, respectively) (Gemma
Reguera, personal communication). This conservation in the family Geobacteraceae is
consistent with regulatory adaptations of these organisms to their unique physiology based on
extracellular electron transfer, as previously demonstrated for other sigma factors (57).
RpoE is not essential for growth, but regulates cell envelope integrity during growth
-

transitions. Colonies of an RpoE mutant strain were easily recovered in medium with acetate
and fumarate with recovery efficiencies comparable to that of other mutants (10). Thus,
although RpoEGsu is the only ECF sigma factor homologue identified in G. sulfurreducens, it was
not essential for growth under these conditions. This contrasts with the essential role of RpoE
-

in E. coli, which prevents the recovery of RpoE mutants (12). However, the growth rates of the
144

-

o

RpoE mutant during exponential growth in NABF medium at 30 C were 0.6-fold lower than in
the wild-type strain (Fig. 5.2A).

-

FIGURE 5.2: RpoE demonstrates impaired growth in NBAF medium compared to WT. (A)
-

o

Growth of the WT (filled circles), and RpoE (open circles) strains at 30 C in NBAF medium. (B-D)
-

Morphology of negatively stained cells of the wild-type (B) and RpoE (C and D) strains by TEM.
-

RpoE mutant cells with cell envelope distortions and damage (C) and abundant spheroplasts
(D) are shown. Bar, 500 nm (B and C), 2 µm (D).

Furthermore, while the wild type resumed exponential growth in NBAF medium upon
-

inoculation, the RpoE mutant had a growth defect (Fig. 5.2A). Extended lag phases (17, 41)
have been reported for RpoE-deficient mutants in other bacteria and have been correlated to
an increased sensitivity of the mutant strains to adapt to even transient growth transitions. This
is because of the extensive reprogramming of the cell’s physiology that is required to adapt to
145

the new nutritional status and resume exponential growth. Such nutritional stress often leads
to periplasmic shock and the activation of the RpoE-mediated response (8).
-

Although we did not observe any defect in the transition of the RpoE cells from
exponential to stationary phase (Fig. 5.2A), TEM micrographs of negatively-stained cells of the
-

wild-type and RpoE mutant were consistent with losses of cell envelope integrity in the mutant
(Fig. 5.2B-D). In contrast to the smooth appearance of wild-type cells (Fig. 5.2B), mutant cells
from cultures transitioning from exponential to stationary phase had outer membrane
distortions and damage (Fig. 5.2C). Periplasmic shock during growth transitions can severely
damage the cell envelope and result in morphological changes such as those noted for the
-

RpoE cells (41). Furthermore, the mutant cultures contained abundant spheroplasts (Fig. 5.2D).
Depletion of periplasmic contents due to the formation of spheroplasts has been proposed to
mimic the effect of misfolded proteins caused by extracytoplasmic stressors, which in E. coli
induces the Cpx cell envelope stress response (61).
Interestingly, attempts to complement the rpoE mutation in trans using a medium-copy
-

plasmid were unsuccessful. Control reactions demonstrated that RpoE can survive
electroporation, as both controls lacking foreign DNA grew as confluent lawns when plated.
However, with the introduction of foreign DNA (pRG5 or pRG5::rpoE) no growth was observed.
This effect was reproducible, as the same result was repeated using several preparations of
fresh competent cells, and numerous electroporation attempts. Thus, we concluded that

146

-

electrocompetent RpoE cells are unable to recover after the introduction of foreign DNA, likely
due to the lack of cell envelope integrity, as described above.
Another growth transition that Geobacter spp. encounter in the environment and that is
of interest for applications in bioremediation and microbial fuel cells is the transition from the
planktonic to the biofilm mode of growth. Biofilm formation requires proper functioning of the
bacterial cell envelope to ensure surface colonization as well as adaptation to growth on the
surface. To study a potential role for RpoE during biofilm formation in G. sulfurreducens, we
-

compared the biofilm-forming abilities of the wild-type and RpoE mutant strains on glass
surfaces under conditions in which the wild type predominatly grows as a planktonic culture
-

(Fig. 5.3A). In contrast to the wild type, the RpoE mutant predominantly grew as a biofilm.

147

FIGURE 5.3: Regulation of biofilm formation by RpoE. (A) Planktonic (circles) and biofilm
-

(squares) growth of the wild-type (solid symbols) and RpoE mutant (open symbols) strains. (B
and C) CSLM micrographs showing top (top panels) and side (bottom panels) projections of 48 h
-

biofilms of the wild-type (B) and RpoE (C) strains on glass coverslips. Cells were stained with
the BacLight viability dyes (green, live cells; red, dead cells). Bar, 20 µm.

After 48 h, when the wild-type biofilms were still thin and unstructured (Fig. 5.3B), the
-

RpoE biofilms had the highest levels of biofilm biomass (Fig. 5.3C). Thus, although inactivation
-

of RpoE in other bacteria impairs biofilm growth (75) the RpoE mutant of G. sulfurreducens
promotes it. It is interesting to note that the biofilm physiology is thought to be analogous to
that of cells entering stationary phase (19). The morphological changes noted for the RpoE

-

cells of G. sulfurreducens in early stationary phase (Fig. 5.2C) are likely to change the physical
148

and chemical properties of the cell surface in a way that could promote adhesion to glass
surfaces. Increased viability in stationary phase has also been reported for RpoE-deficient
mutants in other bacteria (30, 56). In E. coli, for example, the expression of the rpoE gene
promotes cell lysis in early stationary phase but not in exponential phase (30). Hence, RpoEGsu
could mediate similar functions and prevent cell lysis during biofilm formation, thus promoting
biofilm growth.
RpoE is required for stress tolerance. In E. coli, the RpoE transcriptional cascade plays a
major role is responding to stressful environmental conditions that are caused by any internal
or external parameter that affects protein folding and stability in the cell envelope, and
increases the accumulation of misfolded or unfolded proteins in the periplasm (50, 52). RpoE
then activates the genes that are required to degrade and/or repair the misfolded proteins and
synthesize new ones, when appropriate. Protein misfolding in the bacterial periplasm is
counteracted by the activation by RpoE of genes encoding periplasmic protein-folding catalysts
and degradation factors as well as secretion of newly synthesized proteins to replace those
damaged. Consistent with this repair mechanism, genes involved in the degradation and repair
of proteins, protein export via the Sec apparatus, amino acid transport, and protein synthesis
were found to be activated by RpoEGsu (Gemma Reguera, personal communication). Similarly,
genes involved the synthesis of cell envelope components, such as lipoproteins, membrane
proteins, and fatty acid and phospholipids metabolism, as well as genes involved in
gluconeogenesis, biosynthesis and anaplerosis, were also activated by RpoEGsu (Gemma

149

Reguera, personal communication), also indicating a role for RpoEGsu in the repair mechanisms
that restore and replace damaged components of the cell envelope.
Hence, we investigated the effect of extracytoplasmic stressors commonly found in
environments inhabited by Geobacter bacteria such as O2 intrusions, pH, and temperature
-

shifts on the growth of the RpoE mutant in comparison with the wild type (Fig. 5.4).

150

-

FIGURE 5.4: The effect of environmental stressors on the growth of the wild-type (solid symbols) and the RpoE mutant (open
symbols) strains. Stressors include O2 exposure (A), pH (B) and temperature (C). Each data point represents the result of triplicate
-

biological replicates. Inset in (A) shows cuvettes of WT (left) and RpoE (right) following exposure to oxygen.

151

Oxygen intrusions are common in the subsurface environments inhabited by Geobacter
bacteria. Not surprisingly, G. sulfurreducens, though first described as a strict anaerobe (6), can
tolerate long-term exposure to O2 and grow with O2 under microaerophilic conditions for
limited periods of time (39). Survival in the presence of O2 was significantly inhibited in the
-

RpoE mutant compared to the wild type and cell lysis was apparent after just 24 h of exposure
to air (Fig. 5.4A, inset). This is consistent with a role of RpoEGsu in the oxidative stress response
in G. sulfurreducens. Previous studies (57) also demonstrated a role for the stationary-phase
sigma factor RpoS, which also controls the expression of RpoE, in aerotolerance in G.
sulfurreducens. Thus, RpoE and RpoS likely cooperate to regulate the response to oxidative
stress in this organism. Two stress responses that are not mediated by RpoSGsu are resistance
to acidic pH (pH 6 for 60 min) and high temperature (45°C for 7 days) (57). Similarly, RpoEGsu
had no apparent function in resistance to acidic pH (pH 6 for 48 h). However, it was involved in
growth survival after exposure for 24 h to basic pH (pH 9) (Fig. 5.3B). Inasmuch as Geobacter
spp. use acids as electron donors (e.g., acetate) or acceptors (e.g., fumarate), this result is
consistent with an environmental adaptation to the low pHs inhabited by these organisms.
-

The RpoE mutant was also sensitive to low, suboptimal growth temperatures, yet
tolerated high temperatures better than the wild type, having optimal growth temperatures at
40°C rather than 35°C (Fig. 5.4C). This heat-tolerant phenotype can be explained by three
cytoplasmic small heat shock proteins (Hsp20-family) that are normally repressed by RpoEGsu,

152

-

but are overexpressed in the RpoE mutant (Gemma Reguera, personal communication). Small
heat shock proteins associate with cytoplasmic denaturated proteins and protein aggregates,
which are produced during exposure to stressors, and facilitate chaperone-mediated
disaggregation and refolding (14, 18, 38, 53, 54). The activation of genes encoding cytoplasmic
small heat shock proteins thus serves as an indicator of the cytoplasmic stress level in the cells.
Their negative regulation by RpoEGsu is in agreement with a shift of the stress response to the
-

cell envelope, as well as the observed heat-tolerant phenotype of the RpoE mutant. Analyses
of rpoE transcripts by RT-PCR confirmed the inverse correlation between rpoE expression and
temperature (Fig. 5.5), thereby supporting the involvement of RpoE in growth at low
suboptimal temperatures that are relevant to the environmental survival of Geobacter bacteria
in the subsurface.

FIGURE 5.5: Reverse Transcriptase (RT)-PCR of rpoE and recA transcripts. Transcripts from WT
-

cells grown at 25°C, 30°C or 35°C or RpoE cells grown at 30°C are included. Controls with
(+RT) or without (-RT) reverse transcriptase are shown for the rpoE transcripts as well as
controls with WT gDNA.

153

Thus, the mechanisms for coping with cell envelope damage by stressors commonly
found in environments inhabited by Geobacter bacteria are, at least partially, RpoE-dependent.
RpoE regulates electron transfer at the cell envelope. As the integrity of the cell
envelope is required for extracellular transfer reactions and cell viability in G. sulfurreducens
(9), we investigated the ability of resting cell suspensions of the wild-type and RpoE-deficient
mutant strains to catalyze the reduction of soluble (Fe(III) citrate) and insoluble (Fe(III) oxides)
forms of Fe(III) as electron acceptors with acetate or H2 serving as electron donors and (Fig.
5.6).
0.6

A

2.4

mmol.mg-1.min-1

mmol.mg-1.min-1

3.2

1.6

0.8

0

B

0.4

0.2

0
Acetate

Hydrogen

Acetate

Hydrogen
-

FIGURE 5.6: Fe(III) reductase activity of resting cell suspensions. WT (filled blocks) and RpoE
(open blocks) strains were tested using soluble (Fe(III) citrate) (A) or insoluble (poorly crystalline
Fe(III) hydroxide) (B) as electron acceptors.

As shown in Fig. 5.6A, the reduction of Fe(III) citrate was similar in the two strains when
-

acetate was the electron donor. However, the Fe(III) citrate reduction activity of the RpoE cells

154

was only 12% ( 4%) of wild-type activity when H2 was used as an electron donor (Fig. 5.6A).
Acetate is oxidized in the cytoplasm, whereas H2 is oxidized in the periplasm. Thus, the defects
in the reduction of Fe(III) citrate when H2 was the electron donor are consistent with a reduced
-

activity in H2 oxidation. In support of this, microarray analyses of the RpoE mutant in reference
to the wild-type strain revealed a decrease in transcription of genes in the hybSABLP operon in
the mutant (Gemma Reguera, personal communication), which encodes the respiratory
hydrogenase of G. sulfurreducens (11). The Hyb hydrogenase is essential for H2-dependent
growth in G. sulfurreducens (11) and, as a result, the diminished transcript levels for the hyb
-

-

genes in the RpoE mutant also correlate with reduced uptake hydrogenase activity in the RpoE
mutant (51% (± 3%)) of the activity measured in resting cell suspensions of the wild-type cells
(Gemma Reguera, personal communication).
-

By contrast, the RpoE mutant was defective at the reduction of insoluble Fe(III) oxides
with both acetate and H2 (23  5% and 20  7% of wild-type activities with acetate and H2,
respectively) (Fig. 5.6B). Although acetate oxidation takes place in the cytoplasm, the electrons
generated in this intracellular reaction must be transferred to periplasmic and then to outer
membrane c-cytochromes (5, 32, 33, 36, 40, 47) and conductive pili (63) to reduce the insoluble
Fe(III) oxides present outside the cell. Transcript levels for numerous c-type cytochromes were
diminished or up-regulated in the RpoE-deficient mutant (Gemma Reguera, personal
communication). C-cytochromes are key components of the electron transport chain that takes

155

the electrons across the periplasmic space and to the outer membrane in Geobacter cells (20,
32, 33, 36, 40, 68). Among the c-cytochrome genes that were activated by RpoE were those
encoding three outer-membrane cytochromes important for Fe(III) reduction, such as OmcS,
OmcD, and OmcG, and genes encoding proteins required for c-cytochrome biogenesis. OmcS is
required for the reduction of insoluble Fe(III) oxides (47) and electrodes (24) by G.
sulfurreducens while OmcD has been proposed to play a compensatory role during adaptation
to electron transfer disruption (35). Defects in OmcG have been linked to decreased rates of
Fe(III) reduction through an indirect effect on the levels of key c-cytochromes such as OmcB
(33). Protein misfolding in the periplasm also affects the integrity of proteins involved in the
assembly of extracellular appendages spanning the bacterial envelope such as pili (26, 29, 55). .
In fact, genes required for type IV pilus biogenesis (GSU2030 and GSU2031) were down-

regulated in the RpoE mutant (Gemma Reguera, personal communication). The pili of G.
sulfurreducens are conductive protein appendages that function as nanowires to transfer
electrons from the cell envelope to insoluble Fe(III) oxides, their natural electron acceptor for
growth (63), and to uranium (9). Thus, stresses affecting the integrity of the cell envelope also
are likely to affect the assembly of G. sulfurreducens pilus nanowires and their ability to
function as electronic conduits to extracellular electron acceptors.
Implications. This first study of RpoE in a member of the Geobacteraceae demonstrates
the functional conservation of a global master regulator for cell envelope integrity and
homeostasis and highlights specific adaptations to the natural environments inhabited by these
bacteria. As in other bacteria, RpoE responded to extracytoplasmic stressors that lead to cell
envelope damage and protein misfolding in the periplasm and used conserved stress response
156

mechanisms of repair to cope with cell envelope damage. However, the response was specific
to signals relevant to the survival of Geobacter bacteria in the subsurface such as O2 intrusions,
higher pHs and low, suboptimal temperatures. Interestingly, all these stressors lead to slow
growth in G. sulfurreducens, similarly to the slow growth that occurs during the reduction of
Geobacter’s insoluble electron electrons such as Fe(III) oxides or electrodes. Furthermore,
genes required for these reductive processes are part of the RpoE regulon. The adaptation of
the RpoE-mediated response to control processes relevant to survival in the subsurface,
bioremediation of toxic metals and microbial fuel cell performance thus helps explain why G.
sulfurreducens and related organisms function so effectively in Fe(III)-reducing subsurface
environments and in applications that harness their unique reductive activities.

157

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165

CHAPTER 6

CONCLUSIONS AND FUTURE DIRECTIONS

166

The overall objective of this dissertation has been to further the understanding of
bacterial U reduction mechanisms by G. sulfurreducens. In the long-term, this knowledge is
critical to harness microbial U reduction processes so as to develop improved or novel
bioremediation schemes in subsurface environments. In this dissertation I have demonstrated a
novel role for pilus nanowires in U reduction in G. sulfurreducens. Furthermore, the conductive
pili were the primary U reductase in planktonic cells (which is relevant to the physiology of cells
in the groundwater environment) and critical catalytic components in biofilms (a physiological
state relevant to the sediment environment). Additionally, I demonstrated that the U reductive
abilities of the biofilms depended on their stage of development, with maximum reduction
occurring only in the later stages of biofilm development due to the increased production of pili
and redox biofilm matrix as the biofilms grow and mature. Furthermore, I also screened a
library of transposon-insertion mutants for defects in biofilm formation and identified mutants
arrested at various stages of biofilm development and, therefore, defective in attachment,
microcolony formation, and biofilm maturation. Based on the genes that were interrupted by
the transposon in each mutant, I was able to group the mutants in functional categories, thus
enabling the genetic analysis of biofilm development in G. sulfurreducens. Most mutants carried
the transposon insertion in genes annotated in the cell envelope biosynthesis and function
category as well as in electron transport, which is mostly located in the cell envelope. To gain
insights the regulation of the cell envelope in G. sulfurreducens, I characterized a mutant
deficient in the extracytoplasmic function (ECF) sigma factor, RpoE. The results indicate that
this sigma factor is, at least in part, responsible for maintaining the integrity of the cell envelope
in response to environmental stressors such as temperature and oxidative stress.
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Taken together, the results presented in this dissertation provide novel insights into
both the mechanisms of uranium reduction by Geobacter bacteria, and the cellular components
required for biofilm formation and survival in the subsurface. This knowledge will aid in the
development of bioremediation schemes, including the use of biofilms as bioreactive barriers to
immobilize and prevent the spread of U contamination.
Future directions for the work presented in this dissertation include further analysis of
the biofilm-deficient transposon-insertion mutants. Genetic complementation of the pilibiogenesis mutants described in chapter 4 has been carried out by introducing a constitutivelyexpressed WT copy of the interrupted gene in trans. The biofilm formation phenotype of the
complemented strains will allow us to confirm the role of the pili-biogenesis proteins in biofilm
development. Also of special interest is the role that other electron transport proteins
identified in our mutant screen (such as hydrogenases and cytochromes) play in the biofilms.
Future work will include the phenotypic characterization, as well as complementation of these
mutants. Once mutants of interest are characterized in the lab, and selected as potential
molecular markers for biofilm development and/or U reduction in the subsurface, we would
like to take this project to the field to determine the environmental relevance of our genes. The
Zhou lab at the University of Oklahoma has developed functional gene arrays, or “GeoChips”,
which contain probes that target genes involved in a specific process of interest (1). Examples
include genes involved in biogeochemical cycling of organic and inorganic substances,
biodegradation of contaminants, stress response, and antibiotic resistance (1). Geobacter-, pili-,
and biofilm-specific probes could be incorporated in the GeoChip to monitor the activity and
physiological state of Geobacter bacteria in the subsurface during active in situ bioremediation.
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Thus, genes encoding proteins involved in U reduction, such as components of the pilus
apparatus, or those expressed during biofilm development could provide molecular markers to
assess the effectiveness of the bioremediation scheme and enable long-term monitoring. This
will provide valuable information essential to the development of successful in situ
bioremediation schemes.
Additionally, the identification of specific cellular components responsible for U
reduction in Geobacter bacteria will enable the manipulation of their expression or activity to
promote increased removal and reduction of U. One possibility would be to increase the
biological expression of the pilus nanowires. This could be accomplished using genetic
engineering to develop hyperpiliated strains that also constitutively express the nanowires.
Alternatively, and to avoid limitations related to the introduction of genetically engineered
strains into the environment, adaptive evolution could be used to isolate hyperpiliated strains.
The evolution experiments could involve serial transfers of cultures grown under the pressure
to produce pili such as suboptimal temperatures (2), growth with Fe(III) oxides as sole electron
acceptor (2), and biofilm formation (3). These strains could be used to bioremediate
environments where Geobacter bacteria are not present or where metal reduction cannot be
stimulated in situ. Furthermore, the pilus nanowires could also be mass-produced and
assembled in vitro to develop man-made devices for the bioremediation of U in environments
that prevent the growth of microorganisms, such as high-radiation environments. Such
biomimetic interfaces could integrate not only the pilus nanowires but also other U reductase
components identified in my work, such as outer membrane cytochromes, which could also be
produced in vitro and immobilized on the surface of an electrode. This would create a synthetic
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and biodegradable mimic of the Geobacter cell envelope that could be used to safely remove
and precipitate U in environments inhospitable to microbial growth. It could also easily be
removed from the subsurface, thereby also removing the U and preventing further
contamination, while not requiring large-scale excavation of the site. Our lab is currently
involved in the development of these devices in collaboration with Dr. Mark Worden’s lab in
Chemical Engineering (Michigan State University).
In addition to the manipulation of pili production in planktonic cells described above,
similar methods could be used for the development of biofilm biobarriers with increased pili
and biomass so as to enhanced U reductase activity and enable the prolongued reduction of U
and tolerance to higher concentrations of the contaminants. Additionally, we can use the
molecular markers we have identified to monitor the U reductase activity of the biofilms, which
would allow us to better predict the fate of U in the environment, and contribute to the longterm stewardship of contaminated sites.

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REFERENCES

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REFERENCES

1.

He, Z., J. D. Van Nostrand, and J. Zhou. 2012. Applications of functional gene
microarrays for profiling microbial communities. Current opinion in biotechnology
23:460-466.

2.

Reguera, G., K. D. McCarthy, T. Mehta, J. S. Nicoll, M. T. Tuominen, and D. R. Lovley.
2005. Extracellular electron transfer via microbial nanowires. Nature 435:1098-1101.

3.

Reguera, G., R. B. Pollina, J. S. Nicoll, and D. R. Lovley. 2007. Possible Nonconductive
Role of Geobacter sulfurreducens Pilus Nanowires in Biofilm Formation. J. Bacteriol.
189:2125-2127.

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