Investigation of electron transfer mechanisms
in electrochemically active microbial biofilms
Von der Fakultät für Lebenswissenschaften
der Technischen Universität Carolo-Wilhelmina
zu Braunschweig
zur Erlangung des Grades eines
Doktors der Naturwissenschaften
(Dr. rer. nat.)
genehmigte
D i s s e r t a t i o n
Kumulative Arbeit
von Alessandro Alfredo Carmona Martínez
aus Oaxaca / Mexiko
1. Referentin oder Referent: Prof. Dr. Uwe Schröder
2. Referentin oder Referent: Prof. Dr. Rainer Meckenstock
eingereicht am: 30.05.2012
mündliche Prüfung (Disputation) am: 05.10.2012
Druckjahr 2012
Vorveröffentlichungen der Dissertation
Teilergebnisse aus dieser Arbeit wurden mit Genehmigung der Fakultät für
Lebenswissenschaften, vertreten durch den Mentor der Arbeit, in folgenden Beiträgen vorab
veröffentlicht:
Publikationen
Chapter 2: A.A. Carmona-Martinez, F. Harnisch*, L.A. Fitzgerald, J.C. Biffinger, B.R.
Ringeisen, U. Schröder, Cyclic voltammetric analysis of the electron transfer of Shewanella
oneidensis MR-1 and nanofilament and cytochrome knock-out mutants, Bioelectrochemistry,
81 (2011) 74-80.
Chapter 3: A.A. Carmona-Martínez, F. Harnisch*, U. Kuhlicke, T.R. Neu, Uwe Schröder,
Electron transfer and biofilm formation of Shewanella putrefaciens as function of anode
potential, Bioelectrochemistry, (2012) Accepted.
Chapter 4: A.A. Carmona-Martinez, K.H. Ly, P. Hildebrandt, U. Schröder, F. Harnisch*, D.
Millo*, Spectroelectrochemical analysis of intact microbial biofilms of Shewanella species for
sustainable energy production, In preparation, (2012).
Chapter 5: S. Chen, H. Hou, F. Harnisch, S. A. Patil, A. A. Carmona-Martínez, S. Agarwal,
Y. Zhang, S. Sinha-Ray, A. L. Yarin*, A. Greiner*, U. Schröder*, Electrospun and solution
blown three-dimensional carbon fiber nonwovens for application as electrodes in microbial
fuel cells, Energy & Environmental Science, 4 (2011) 1417-1421.
Chapter 6: S. Chen, G. He, A.A. Carmona-Martínez, S. Agarwal, A. Greiner, H. Hou*, U.
Schröder*, Electrospun carbon fiber mat with layered architecture for anode in microbial fuel
cells, Electrochemistry Communications, 13 (2011) 1026–1029.
Chapter 7: S.A. Patil, F. Harnisch*, C. Koch, T. Hübschmann, I. Fetzer, A.A. Carmona-
Martínez, S. Müller*, U. Schröder, Electroactive mixed culture derived biofilms in microbial
bioelectrochemical systems: the role of pH on biofilm formation, performance and
composition, Bioresource Technology, 102 (2011) 9683–9690.
Chapter 8: F. Harnisch*, C. Koch, I, Fetzer, A. A. Carmona-Martínez, S. F. Hong, S. A.
Patil, T. Hübschman, U. Schröder, S. Müller*, Electroactive mixed culture derived biofilms in
microbial bioelectrochemical systems: the role of inoculum and substrate on biofilm
formation and performance, In preparation (2012).
*indicates authors of correspondence
Tagungsbeiträge
Oral presentations:
A.A. Carmona-Martínez, F. Harnisch, U. Kuhlicke, T.R. Neu, U. Schröder. 2012. Electron
Transfer and Biofilm Formation of Shewanella putrefaciens as Function of Anode Potential.
Submitted for oral presentation at the EU-ISMET meeting: From extracellular electron
transfer to innovative process development, Ghent (Belgium), September 27th – 28th, 2012.
A.A. Carmona-Martínez, 2009. Microbial fuel cells: an alternative for the production of
clean electricity. Abstract F128. Presented at the German Academic Exchange Service
Scholarship Holders Meeting. Hanover (Germany). June 19th – 21th, 2009.
Poster presentations:
A.A. Carmona-Martínez, S. Patil, F. Harnisch, U. Schröder, S. Chen, C. Greiner, A.
Agarwal, H. Hou, Y. Zhang, S. Sinha-Ray, A. Yarin. 2011. High Surface Area Electrospun
and Solution-blown Carbonized Nonwovens to Enhance the Current Density in
Bioelectrochemical Systems (BES). Abstract ELE 026. Presented at Wissenschaftsforum
Chemie 2011, Bremen (Germany), September 4th – 7th, 2011.
A.A. Carmona-Martínez, F. Harnisch, U. Schröder. 2010. Analysis of the electron transfer
and current production of Shewanella oneidensis MR-1 wild-type and derived mutants.
Abstract P058. Presented at Electrochemistry 2010: From microscopic understanding to
global impact, Bochum (Germany), September 13th – 15th, 2010.
A.A. Carmona-Martínez, F. Harnisch, U. Schröder. 2009. Cyclic voltammetry as a useful
technique to characterize electrochemically active microorganisms: Shewanella putrefaciens.
Abstract AE15. Presented at Wissenschaftsforum Chemie 2009, Frankfurt am Main
(Germany), August 30th – September 2nd, 2009. ISBN: 978-3-936028-59-1.
„Gedruckt mit Unterstützung des Deutschen Akademischen
Austauschdienstes“
To Yolanda, Jesús and Virginia,
for their love and support...
Acknowledgements
First and foremost, I express my gratitude towards my supervisor Prof. Dr. Uwe Schröder for
supporting me since the very first moment I applied for the scholarship to conduct Ph.D.
studies in Germany. Prof. Schröder encouraged me to pursue my own ideas while providing
me invaluable academic freedom and substantial support throughout my entire Ph.D.
I would like to thank as well Dr. Falk Harnisch for his supervision, critical suggestions and
academic inspiration. I want also to thank all the time he has invested in my thesis with
constant guidance during design, planning, data analysis and manuscript writing.
I deeply appreciate the financial and logistic support by the German Academic Exchange
Service providing me a Ph.D. scholarship that allowed me not only to conduct my thesis work
but also by procuring all necessary support to enjoy the academic German culture.
Furthermore, I thank the financial support by the Mexican Secretariat of Public Education for
providing me a complementary Ph.D. scholarship during my stay in Germany.
I am very much grateful to Dr. Sunil A. Patil and Dr. Siang-Fu Hong for valuable
experimental assistance, cooperation and fun time during my stay at the Technischen
Universität Carolo-Wilhelmina zu Braunschweig. Thanks to their hands-on experience, I was
able to solve in a successful way several experimental obstacles.
I would like to sincerely acknowledge the following people for their support and successful
collaboration: 1) Dr. B.R. Ringeisen, Dr. L.A. Fitzgerald and Dr. J.C. Biffinger at the Naval
Research Laboratory in Washington, USA; 2) Dr. T.R. Neu and Ute Kuhlicke at the
Helmholtz Centre in Magdeburg, Germany; and finally 3) Dr. D. Millo, K.H. Ly and Prof. Dr.
P. Hildebrandt at the TU Berlin.
I thank all former and current members of the Sustainable Chemistry and Energy Research
group at the TU Braunschweig for their individual contributions to a very friendly research
atmosphere full of respect and kind collaboration with its invaluable 10 am coffee break
together with the social activities in the group, key components of an enjoyable research.
I express my gratefulness towards my friend circle in Braunschweig.
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Table of contents (brief)
Chapter 1 Extracellular electron transfer in Bioelectrochemical systems: bridge between
natural environments and applied technologies...................................................1
Part I Electron transfer mechanisms of pure culture biofilms of
Shewanella spp.
Chapter 2 Cyclic voltammetric analysis of the electron transfer of Shewanella oneidensis
MR-1 and nanofilament and cytochrome knock-out mutants...........................33
Chapter 3 Study of Shewanella putrefaciens biofilms grown at different applied potentials
using cyclic voltammetry and confocal laser scanning microscopy..................47
Chapter 4 Spectroelectrochemical analysis of intact microbial biofilms of Shewanella
putrefaciens for sustainable energy production.................................................61
Part II Porous 3D carbon as anode materials for performance of
electrochemically active mixed culture biofilms
Chapter 5 Electrospun and solution blown three-dimensional carbon fiber nonwovens for
application as electrodes in microbial fuel cells................................................71
Chapter 6 Electrospun carbon fiber mat with layered architecture for anode in microbial
fuel cells.............................................……………………………....................82
Part III The influence of external factors on electrochemically active
mixed culture biofilms
Chapter 7 Electroactive mixed culture derived biofilms in microbial bioelectrochemical
systems: the role of pH on biofilm formation, performance and
composition.......................................................................................................90
Chapter 8 Electroactive mixed culture derived biofilms in microbial bioelectrochemical
systems: the role of inoculum and substrate on biofilm formation and
performance.....................................................................................................108
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Table of contents (extended)
1 Extracellular electron transfer in Bioelectrochemical systems: bridge between
natural environments and applied technologies .......................................................................... 1
1.4.1.1 DET via membrane-bound redox-enzymes .............................................................................. 5
1.4.1.1.1 Shewanella oneidensis DET via membrane-bound redox-enzymes .................................... 5
1.4.1.1.2 Geobacter sulfurreducens DET via membrane-bound redox-enzymes ............................... 6
1.4.1.2 DET via self-produced microbial nanowires ............................................................................ 6
1.4.1.2.1 Geobacter sulfurreducens DET via self-produced microbial nanowires ............................. 7
1.4.1.2.2 Shewanella oneidensis DET via self-produced microbial nanowires .................................. 7
1.4.2.1 MET via artificial exogenous mediator molecules ................................................................... 9
1.4.2.2 MET via natural exogenous mediator molecules ..................................................................... 9
1.4.2.3 MET via self-produced mediator molecules ............................................................................. 9
1.5.1.1 Microbial fuel cells ..................................................................................................................15
1.5.1.2 Microbial electrolysis cells ......................................................................................................15
1.5.1.3 Microbial desalination cells .....................................................................................................15
1.5.1.4 Microbial solar cells ................................................................................................................16
1.5.1.5 Enzymatic fuel cells ................................................................................................................16
1.1 Prelude ................................................................................................................................................... 1
1.2 Ecological significance of insoluble metal electron acceptors: the example of iron ............................. 2
1.3 Electron transfer processes in the environment ..................................................................................... 3
1.4 Microbial extracellular electron transfer mechanisms ........................................................................... 4
1.4.1 Microbial direct extracellular electron transfer (DET) ...................................................................... 5
1.4.2 Microbial mediated extracellular electron transfer (MET) ................................................................ 8
1.5 Bioelectrochemical systems (BESs) .....................................................................................................11
1.5.1 Types of Bioelectrochemical systems ..............................................................................................13
1.6 Performance of Bioelectrochemical systems ........................................................................................16
1.6.1 Performance based on the improvement of electrode materials .......................................................18
1.6.2 Performance based on the study of environmental factors affecting biofilm formation ..................19
1.7 Aim of this Dissertation ........................................................................................................................21
1.8 Structure of the Thesis and personal contribution ................................................................................22
1.9 Comprehensive summary .....................................................................................................................26
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2 Cyclic voltammetric analysis of the electron transfer of Shewanella oneidensis MR-1
and nanofilament and cytochrome knock-out mutants ...................................................... 33
2.1.1.1 Direct electron transfer (DET) .................................................................................................34
2.1.1.2 Mediated electron transfer (MET) ...........................................................................................36
3 Study of Shewanella putrefaciens biofilms grown at different applied potentials
using cyclic voltammetry and confocal laser scanning microscopy.. ................................. 47
2.1 Introduction ..........................................................................................................................................33
2.1.1 Extracellular electron transfer mechanisms of S. oneidensis MR-1 wild type and mutants .............34
2.2 Materials and methods ..........................................................................................................................36
2.2.1 General conditions ...........................................................................................................................36
2.2.2 Cell cultures and media ....................................................................................................................36
2.2.3 Bioelectrochemical experiments ......................................................................................................37
2.2.4 Data processing ................................................................................................................................37
2.3 Results and discussion ..........................................................................................................................38
2.3.1 Bioelectrochemical current production ............................................................................................38
2.3.2 Cyclic voltammetric analysis and data processing ...........................................................................39
2.4 Conclusions ..........................................................................................................................................46
3.1 Introduction ..........................................................................................................................................47
3.1.1 Influence of the electrode potential on electroactive microbial biofilms .........................................49
3.2 Materials and methods ..........................................................................................................................50
3.2.1 General conditions ...........................................................................................................................50
3.2.2 Cell cultures and media ....................................................................................................................50
3.2.3 Bioelectrochemical set-up and experiments .....................................................................................51
3.2.4 Electrochemical data processing ......................................................................................................51
3.2.5 Confocal Laser Scanning Microscopy..............................................................................................52
3.3 Results and discussion ..........................................................................................................................52
3.3.1 Bioelectrochemical current production ............................................................................................52
3.3.2 Cyclic voltammetric analysis ...........................................................................................................54
3.3.3 Biofilm imaging using confocal laser scanning microscopy (CLSM) .............................................58
3.4 Conclusions ..........................................................................................................................................60
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4 Spectroelectrochemical analysis of intact microbial biofilms of Shewanella
putrefaciens for sustainable energy production ................................................................... 61
5 Electrospun and solution blown three-dimensional carbon fiber nonwovens for
application as electrodes in microbial fuel cells ................................................................... 71
5.2.1.1 Gas-assisted electrospinning carbon fiber mat (GES-CFM) ...................................................73
5.2.1.2 Electrospun carbon fiber mat (ES-CFM) .................................................................................74
5.2.1.3 Solution-blown carbon fiber mat (SB-CFM) ...........................................................................74
4.1 Introduction ..........................................................................................................................................61
4.2 Materials and methods ..........................................................................................................................64
4.2.1 Materials and methods .....................................................................................................................64
4.2.2 Cell cultures and media ....................................................................................................................64
4.2.3 Electrochemical set-up for the growth of anodic electrocatalytic biofilms ......................................65
4.2.4 Growth of anodic electrocatalytic biofilms ......................................................................................66
4.2.5 Cyclic voltammetry ..........................................................................................................................66
4.2.6 Electrochemical data processing ......................................................................................................66
4.2.7 Spectroelectrochemical set-up for SERRS measurements ...............................................................66
4.2.8 SERRS measurements ......................................................................................................................66
4.3 Results and discussion ..........................................................................................................................67
4.3.1 Bioelectrochemical current production ............................................................................................67
4.4 Conclusions ..........................................................................................................................................70
5.1 Introduction ..........................................................................................................................................71
5.2 Materials and methods ..........................................................................................................................73
5.2.1 Carbon fiber preparation ..................................................................................................................73
5.2.2 Electrode preparation .......................................................................................................................75
5.2.3 Bioelectrochemical experiments ......................................................................................................75
5.3 Results and discussion ..........................................................................................................................76
5.3.1 Biocatalytic current generation at modified carbon electrodes ........................................................76
5.3.2 Analysis of electroactive biofilms grown at modified carbon electrodes with Scanning electron
microscopy ....................................................................................................................................................77
5.3.3 Cyclic voltammetry of electroactive biofilms grown at modified carbon electrodes .......................79
5.4 Conclusions ..........................................................................................................................................81
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6 Electrospun carbon fiber mat with layered architecture for anode in microbial fuel
cells...........................................................................................................................................82
7 Electroactive mixed culture derived biofilms in microbial bioelectrochemical
systems: the role of pH on biofilm formation, performance and composition ................. 90
7.2.8.1 Flow-cytometry .......................................................................................................................94
7.2.8.1.1 Sample fixation and DNA staining .................................................................................... 94
7.2.8.1.2 Multiparametric flow-cytometry ........................................................................................ 94
7.2.8.2 T-RFLP and Sequencing .........................................................................................................95
6.1 Introduction ..........................................................................................................................................82
6.2 Materials and methods ..........................................................................................................................83
6.2.1 Carbon fiber preparation ..................................................................................................................83
6.2.2 Electrode preparation .......................................................................................................................84
6.2.3 Bioelectrochemical measurements ...................................................................................................84
6.2.4 SEM imaging ...................................................................................................................................84
6.3 Results and discussion ..........................................................................................................................85
6.3.1 Properties and performance of carbon fiber mat electrode materials ...............................................85
6.3.2 Biocatalytic current generation at carbon fiber mat electrode materials ..........................................87
6.3.3 Analysis of electroactive biofilms grown at carbon fiber mat electrode materials with Scanning
electron microscopy ......................................................................................................................................87
6.4 Conclusions ..........................................................................................................................................89
7.1 Introduction ..........................................................................................................................................90
7.2 Materials and methods ..........................................................................................................................91
7.2.1 General conditions ...........................................................................................................................91
7.2.2 Electrochemical set-up .....................................................................................................................92
7.2.3 Microbial inoculum and growth medium .........................................................................................92
7.2.4 Biofilm growth (fed-batch experiments) ..........................................................................................92
7.2.5 Biomass determination .....................................................................................................................93
7.2.6 Metabolic analysis ............................................................................................................................93
7.2.7 Continuous flow mode operation and pH-regime studies ................................................................93
7.2.8 Microbiological analysis ..................................................................................................................94
7.3 Results and discussion ..........................................................................................................................96
7.3.1 Biofilm formation and performance at different constant pH ..........................................................96
7.3.2 Biofilm performance at varying pH-environment during operation .................................................97
7.3.3 Influence of the pH and buffer capacity on the electron transfer .....................................................99
7.3.4 Microbial biofilm analysis .............................................................................................................101
7.4 Conclusions ........................................................................................................................................107
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8 Electroactive mixed culture derived biofilms in microbial bioelectrochemical
systems: the role of inoculum and substrate on biofilm formation and performance ... 108
9 Supplementary information: Chapter II .................................................................... 120
10 Supplementary information: Chapter III ................................................................... 130
11 Supplementary information: Chapter VII ................................................................. 136
12 References ...................................................................................................................... 148
8.1 Introduction ........................................................................................................................................108
8.2 Materials and methods ........................................................................................................................111
8.2.1 General conditions .........................................................................................................................111
8.2.2 Electrochemical set-up ...................................................................................................................111
8.2.3 Microbial inoculum and growth medium .......................................................................................112
8.2.4 Biofilm growth in bioelectrochemical half-cells ............................................................................112
8.2.5 Cyclic voltammetry ........................................................................................................................113
8.2.6 Metabolic analysis for coulombic efficiency calculation ...............................................................113
8.3 Results and discussion ........................................................................................................................113
8.3.1 Current density production of enriched microbial electroactive biofilms as a function of microbial
inoculum and substrate ................................................................................................................................113
8.3.2 Bioelectrocatalytic activity of enriched microbial electroactive biofilms as a function of microbial
inoculum and substrate ................................................................................................................................115
8.4 Conclusions ........................................................................................................................................118
11.1 Influence of the buffer capacity ..........................................................................................................136
11.2 Terminal restriction fragment polymorphism (T-RFLP) analysis: Anode biofilm composition at the
different pH values determined by T-RFLP ...................................................................................................137
11.3 Terminal restriction fragment polymorphism analysis: Anode chamber community composition at pH
7 and 9 at different feeding cycles determined by T-RFLP ............................................................................140
11.4 Relationship of community composition when cultivated at different pH and under successive feeding
cycles determined by T-RFLP ........................................................................................................................140
11.5 Flow-cytometric analysis. ...................................................................................................................142
11.5.1 Community structure when cultivated at pH 9 at successive feeding cycles determined by flow
cytometry ....................................................................................................................................................142
11.5.2 Community structure when cultivated at pH 6 at successive feeding cycles determined by flow
cytometry ....................................................................................................................................................143
11.6 Relationship of community structure when cultivated at different pH and under successive feeding
cycles determined by flow cytometry .............................................................................................................144
11.7 Statistical Analysis of flow-cytometric data .......................................................................................145
11.8 Biofilm detachment ............................................................................................................................146
11.9 Multivariate statistical analysis of the flow-cytometric pattern using n-MDS-plots ..........................147
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Index of figures
Figure 1-1 Simplified iron cycle in aquatic environments. Figure drawn with modifications after (Luu and
Ramsay, 2003, Nealson and Saffarini, 1994). ........................................................................................... 3
Figure 1-2 Overall illustration of microbial ET mechanisms found in the literature. A) Direct extracellular
electron transfer via membrane bound cytochromes and conductive nanowires and B) Mediated
extracellular electron transfer via a mediator molecule (Medred
or Medox
) (see text). Here ET
mechanisms are represented in the field of BESs with electrode materials as final electron acceptors but
the same illustration could be applied for bacteria in natural environments using for instance iron oxides
as terminal electron acceptors. Figure drawn with modifications after (Schröder, 2007). ........................ 4
Figure 1-3 Roles of outer membrane cytochromes of A) Shewanella oneidensis and B) Geobacter sulfurreducens
in extracellular electron transfer. IM: inner membrane, OM: outer membrane and PS: periplasm. Figure
drawn with modifications after (Shi, et al., 2009). .................................................................................... 6
Figure 1-4 Scanning electron microscope micrographs of: A) Geobacter sulfurreducens (ATCC 51573)
(Malvankar, et al., 2011); B) Shewanella oneidensis MR-1 (Gorby, et al., 2006); C) Synechocystis sp.
PCC 6803 (Gorby, et al., 2006) and D) co-culture of Pelotomaculum thermopropionicum and
Methanothermobacter thermautotrophicus showing nanowires connecting the two genera (Gorby, et al.,
2006). ........................................................................................................................................................ 8
Figure 1-5 Number of publications reporting the use of “Bioelectrochemical systems” (Scopus data base, January
2012). Illustration based on (Schröder, 2011). ........................................................................................ 12
Figure 1-6 Overall view of Bioelectrochemical systems. Production of electricity and useful metabolites take
place in BESs. These microbial/ enzyme/ organelles based systems consist of an anode (oxidation
process), a cathode (reduction process) and typically a membrane separating both electrodes (see also
Table 1-2). Depending on the membrane specificity (Harnisch and Schröder, 2009), type of catalysts at
both electrodes (Franks, et al., 2010, Rosenbaum, et al., 2011), and the source of the reducing power
(Logan, et al., 2008, Logan, et al., 2006) a diverse spectrum of research and practical applications can
be found (see Section 1.5.1). Drawn with modifications after (Rabaey and Rozendal, 2010). ............... 13
Figure 1-7 Illustration of the enhancement of the anodic current density performance in BESs. Current density
values taken from representative literature data: (Aelterman, et al., 2006, Bond, et al., 2002, Catal, et al.,
2008a, Catal, et al., 2008b, Chen, et al., 2011, Gil, et al., 2003, He, et al., 2011, He, et al., 2005,
Holmes, et al., 2004b, Katuri, et al., 2010, Kim, et al., 1999b, Kim, et al., 1999d, Liu, et al., 2005, Liu,
et al., 2010c, Milliken and May, 2007, Min and Logan, 2004, Park and Zeikus, 2000, Park, et al., 2001,
Torres, et al., 2009, Zhao, et al., 2010b, Zuo, et al., 2006). Illustration based on Ref. (Schröder, 2011).
................................................................................................................................................................ 17
Figure 1-8 Schematic illustration of the research areas within the three chapter I, II and III. .............................. 22
Figure 2-1 Direct (DET) and mediated (MET) electron transfer pathways utilized by S. oneidensis wild type and
mutants. In every scheme it is indicated which strains can perform the respective electron transfer
mechanisms (Chang, et al., 2006, Nielsen, et al., 2010, Rabaey, et al., 2010). A) Electron transfer via the
cytochrome pool. Transmembrane pilus electron transfer via B) pil-type pilus and via C) msh-type
pilus, and D) biofilm formation behaviour. OM: Outer membrane and IM: Inner membrane................ 35
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Figure 2-2 A) and B) CVs for non-turnover conditions for S. oneidensis WT and mutants using a scan rate of 1
mV s−1
; C and D) provide the respective baseline corrected curves. ...................................................... 39
Figure 2-3 A) and B) CVs for turnover conditions for S. oneidensis WT and mutants using a scan rate of 1 mV
s−1
. ........................................................................................................................................................... 40
Figure 2-4 Plot of the base line corrected height of the oxidation peak of redox-system I (Δi−0.2) as function of
the maximum chronoamperometric current density of the respective microbial culture. ....................... 42
Figure 2-5 Plot of the corrected turnover CV signal and the performed analysis on the example of S. oneidensis
MR-1. (Similar plots of all strains can be found in Fig. S9-8 and Fig. S9-9 in the Supplementary
Information for Chapter 2). ..................................................................................................................... 43
Figure 3-1 Representative chronoamperometric fed-batch cycles of S. putrefaciens at graphite electrodes; applied
potentials: -0.1, 0, +0.1, +0.2, +0.3 and +0.4 V vs. Ag/AgCl; CV measurements during turn-over (A)
and non turn-over (B) conditions respectively. ....................................................................................... 53
Figure 3-2 Chronoamperometric current density of S. putrefaciens as function of the applied electrode potential.
................................................................................................................................................................ 53
Figure 3-3 A) Representative cyclic voltammograms of S. putrefaciens for turn-over conditions and B)
respective first derivatives of the voltammetric curves; scan rate: 1 mV s-1
. .......................................... 55
Figure 3-4 A) Cyclic voltammograms for non turn-over conditions for S. putrefaciens using a scan rate of 1 mV
s−1
; B provides the respective baseline corrected curves. ........................................................................ 56
Figure 3-5 Plot of the base line corrected height (○) and area (□) of the oxidation and reduction peaks of redox-
system shown in Fig. 3-4 as function of the applied potential. For visual convenience, reduction peak
areas are shown as negative values. ........................................................................................................ 57
Figure 3-6 Maximum intensity projection of confocal laser scanning microscopy data sets showing Shewanella
putrefaciens biofilms grown on electrode surfaces at different applied potentials. A) -0.1 V, B) 0 V, C)
+0.1 V, D) +0.2 V, E) +0.3 V and F) +0.4 V; (all vs. Ag/AgCl). Colour allocation: reflection of
electrode – grey, nucleic acid stained bacteria – green. .......................................................................... 58
Figure 3-7 Biofilm quantification of Shewanella putrefaciens biofilms grown on electrode surfaces at different
applied potentials. ................................................................................................................................... 59
Figure 4-1 Principle representation of a BES operating in the DET mode (see below). Electrons derived from the
oxidation of the organic substrate catalyzed by the bacterial cell are shuttled to the electrode via OMCs.
................................................................................................................................................................ 62
Figure 4-2 Electrochemical half cell set-up under potentiostatic control. Insert shows a photograph of the
nanostructured silver ring working electrode. ......................................................................................... 65
Figure 4-3 Chronoamperometric curve of a biofilm formation using a silver ring electrode poised at +0.05 V in a
batch experiment using 18 mM sodium lactate as the substrate and S. putrefaciens cells as biocatalyst.67
Figure 4-4 A) CV of the active biofilm formed on a silver ring electrode under non-turnover conditions (i.e. in
the absence of the substrate sodium lactate) at a scan rate of 1 mV s-1
. B) Respective SOAS baseline
corrected curves. ..................................................................................................................................... 68
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Figure 4-5 SERR spectra of the reduced (upper spectrum) and oxidized (lower spectrum) OMCs, obtained at -
425 and 0 mV, respectively. The spectra were obtained with excitation at λ = 413 nm, laser power of 1
mW, and an acquisition time of 90 s. Potentials refer to the Ag/AgCl (KCl 3 M) reference electrode
(210 mV vs. SHE). .................................................................................................................................. 69
Figure 5-1 (A) Schematic drawing of an electrospinning setup (derived from ref. (Greiner and Wendorff, 2007)).
Solution blowing differs from electrospinning by the use of a high-speed nitrogen jet flow (230–250 m
s-1
) instead of a high voltage electric field to accelerate and stretch the polymer solution into a fibrous
form (Sinha-Ray, et al., 2010). (B) Electrochemical cell for the simultaneous study of different
electrode materials. ................................................................................................................................. 73
Figure 5-2 Biocatalytic current generation at a GES-CFM modified carbon electrode in a model semi-batch
experiment. The GES-CFM electrode was modified by a wastewater-derived secondary biofilm grown
in a half-cell experiment under potentiostatic control. The electrode potential was 0.2 V. .................... 77
Figure 5-3 Scanning electron microscopic images of (A) carbon felt, (B) an electroactive biofilm grown at
carbon felt, (C) GES-CFM, (D) an electroactive biofilm grown at GES-CFM, (E) high resolution image
of GESCFM illustrating the occurrence of inter-fibre junctions, and (F) crosssectional view of GES-
CFM electrode. ....................................................................................................................................... 78
Figure 5-4 Cyclic voltammograms of an electroactive biofilm grown at GESCFM. The voltammograms were
recorded under turnover conditions [in the presence of substrate (10 mM acetate), curve A], as well as
nonturnover conditions (the absence of substrate, curve B). The biofilm was a wastewater-derived
secondary biofilm grown at a potential of 0.2 V under potentiostatic control. The scan rate was 1 mV s-
1. .............................................................................................................................................................. 80
Figure 6-1 A) Top view and B) cross-sectional view SEM images of carbon mat from TP; C) EDX spectra of
NCP-based carbon fiber; D) top view and E) cross-sectional view SEM images of layered-ECFM; F)
cross-sectional view SEM image of 2D-ECFM. ..................................................................................... 86
Figure 6-2 Biocatalytic current generation curves of carbon fiber mats in a half-cell experiment measured at
room temperature. Arrows represent replacement of medium. ............................................................... 87
Figure 6-3 SEM images of biofilms in: A-C belong to layered-CFM; D and E belong to commercial carbon felt;
and F belongs to 2D-ECFM. ................................................................................................................... 88
Figure 7-1 Performance of electroactive biofilms grown and operated at different pH-values: Maximum current
densities (filled circles; derived from chronoamperometric fed-batch experiments at 0.2 V vs. Ag/
AgCl) and coulombic efficiencies (open squares) of primary, wastewater derived biofilms are shown.
The substrate was 10 mM acetate. .......................................................................................................... 96
Figure 7-2 A) Chronoamperometric current density changes (at 0.2 V vs. Ag/ AgCl) for a biofilm initially grown
at pH 7.0 in relation to variations of the growth medium pH (numbers indicate the respective pH-value
of operation); B) Steady state current densities at 0.2 V vs. Ag/ AgCl of biofilms grown at pH 8 (circles)
and pH 7.0 (squares) at varying medium pH (derived from experiments similar as shown in A)). ........ 98
Figure 7-3 Influence of the operational pH: Cyclic voltammograms obtained at different operation pH (using a
constant ionic strength of 50 mM) at a scan rate of 1 mV s-1
during non-turnover conditions for
wastewater derived, acetate-fed biofilm formed at pH 7.0. (For pH 6 to pH 8 steady-state CVs are
shown, for pH 5 the 3rd CV-curve). ..................................................................................................... 100
-x-
Figure 7-4 Bacterial community profiles of the inoculum and the successive media of the anode chamber of a pH
7 grown biofilm (electrode-set 2). The profile of the community is cytometrically determined by the
cells’ DNA content labelled with the A-T specific fluorescent dye DAPI and the cells’ forward scatter
behaviour (FSC). As a result fingerprint-like cytometric patterns emerged as subsets of cells which
gather in numerous clusters of changing cell abundances therein. Up to 250000 cells were analysed and
the dominant sub-populations presented in yellow colour. The peak in the lower left corner of the
histograms represents the noise of the cytometer as well as unstained cell debris. ............................... 103
Figure 7-5 Dalmatian-n-MDS analysis with overlaid cytometric flow-plots derived from anode chamber
communities and anode biofilms when treated over several feeding cycles and different pH-values.
Black patches in flow-plots depict gate positions, cycle number is given with c 1–5 and pH-affiliation
with various grey/black labels (black: pH 7, grey: pH 9, light grey: pH 6, bold fringe around flow-plot:
electrodes; details see text and S11-2 to S11-10 for raw data). ............................................................. 106
Figure 8-1 A) Electrochemical half cell set-up under potentiostatic control and B) Exemplary established
bioelectrochemical active biofilm enriched from primary wastewater fed with acetate. The red color is
mainly caused by the hemes (Jensen, et al., 2010). ............................................................................... 112
Figure 8-2 Bioelectrocatalytic performance of electroactive microbial biofilms derived from different inocula
with fed batch operation in potentiostatically controlled half-cell experiments (+0.2 V vs. Ag/ AgCl) at
graphite rod electrodes. The substrate was 10 mM sodium acetate or sodium lactate respectively. ..... 114
Figure 8-3 Exemplary cyclic voltammograms. Electroactive microbial biofilms derived from different inocula
grown with Sodium acetate (10 mM) recorded during non-turnover (A, C, E and G) and turnover
conditions (B, D, F and H) conditions. The scan rate used was 1 mV s-1
. ............................................ 116
Figure 8-4 Exemplary cyclic voltammograms. Electroactive microbial biofilms derived from different inocula
grown with Sodium lactate (10 mM) recorded during non-turnover (A, C, E and G) and turnover
conditions (B, D, F and H) conditions. The scan rate used was 1 mV s-1
. ............................................ 117
Figure 8-5 Exemplary cyclic voltammograms from electroactive microbial biofilms derived from primary
wastewater grown with 10 mM sodium lactate (A) or 10 mM sodium acetate (B) recorded during
turnover conditions. First derivatives of biofilms grown with sodium lactate (C) or sodium acetate (D).
.............................................................................................................................................................. 118
Figure S9-1 Schematic drawing of an electrochemical cell for the study of the electron transfer mechanisms and
current production. The electrochemical cell consists of an anode, a cathode and, a membrane
separating both. An oxidation process occurs at the anode, in this case lactate oxidation, in which
electrons and protons are produced. The electrons flow to the cathode through an external circuit or
potentiostat in which the electrons can be can be quantified. Meanwhile the protons are released to the
media and lately they migrate to the cathode chamber to react with molecules of water and electrons
finally producing hydrogen for example. Figure drawn with modifications after (Rabaey and Verstraete,
2005, Schröder, 2008). .......................................................................................................................... 121
Figure S9-2 Electrochemical half cell set-up under potentiostatic control. Description: “Top view” shows the 5
necks of the 250 mL flask. In section A-A’ details of the working electrode, counter shielded electrode
and reference electrode are given. In section B-B’ the port for filtrated air, filtrated nitrogen and media
supply are detailed. ............................................................................................................................... 122
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Figure S9-3 Exemplary fed-batch chronoamperometric cycles (0.2 V vs Ag/AgCl) of Shewanella oneidensis
MR-1 Wild-type and knock-out mutants on equally-sized graphite rod anode electrodes, in half cells
utilizing lactate (18 mM) as the electron donor and anodes as electron acceptors. ............................... 123
Figure S9-4 Cyclic voltammetry at 1 mV s-1
(A, C and E) and First derivative plots of CV data (B, D and F) of S.
oneidensis Wild-type (E and F) and mutants (A and B: ΔpilM-Q/ΔmshH-Q; C and D: ΔpilM-Q) during
Turnover conditions. OxT states for oxidation turnover peak, RedT states for reduction turnover peak
and ET states for redox turnover system. Every time 4 exemplary CVs are shown. ............................. 124
Figure S9-5 Continuation of Fig. S9-4. Cyclic voltammetry at 1 mV s-1
(G, I and K) and First derivative plots of
CV data (H, J and L) of S. oneidensis mutants (G and H: ΔmshH-Q; I and J: Δflg; K and L:
ΔmtrC/ΔomcA) during Turnover conditions. OxT states for oxidation turnover peak, RedT states for
reduction turnover peak and ET states for redox turnover system. Every time 4 exemplary CVs are
shown. ................................................................................................................................................... 125
Figure S9-6 Cyclic voltammetry at 1 mV s-1
(A, C and E) and First derivative plots of CV data (B, D and F) of S.
oneidensis Wild-type (E and F) and mutants (A and B: ΔpilM-Q/ΔmshH-Q; C and D: ΔpilM-Q) during
Non-turnover conditions. OxT states for oxidation turnover peak, RedT states for reduction turnover
peak and ET states for redox turnover system. Every time 4 exemplary CVs are shown. ..................... 126
Figure S9-7 Continuation of Fig. S9-6. Cyclic voltammetry at 1 mV s-1
(G, I and K) and First derivative plots of
CV data (H, J and L) of S. oneidensis mutants (G and H: ΔmshH-Q; I and J: Δflg; K and L:
ΔmtrC/ΔomcA) during Non-turnover conditions. OxT states for oxidation turnover peak, RedT states for
reduction turnover peak and ET states for redox turnover system. Every time 4 exemplary CVs are
shown. ................................................................................................................................................... 127
Figure S9-8 Data analysis for each catalytic centre (redox-system I and II). On the left column an exemplary
turnover CV for each strain can be seen. In the center is its respective non-turnover CV. On the right
column the final subtracted CV is presented on which the signal height of each catalytic wave was
estimated at suitable fixed potentials. A-C) ΔpilM-Q/ΔmshH-Q. D-F) ΔpilM-Q. G-I) Wild-type. (see
also Fig. 2-5 in Chapter II for details) ................................................................................................... 128
Figure S9-9 Continuation of Fig. S9-8. Data analysis for each catalytic centre (redox-system I and II). On the left
column an exemplary turnover CV for each strain can be seen. In the center is its respective non-
turnover CV. On the right column the final subtracted CV is presented on which the signal height of
each catalytic wave was estimated at suitable fixed potentials. J-L) ΔmshH-Q. M-N) Δflg, P-R)
ΔmtrC/ΔomcA. (see also Fig. 2-5 in Chapter II for details) ................................................................. 129
Figure S10-1 Electrochemical cell set-up. A) Electrochemical cell hosting six potentiostatic controlled working
electrodes without S. putrefaciens cells. B) Electrochemical cell with M1 growth media inoculated with
whole cells of S. putrefaciens. Insert: photograph showing a reddish pellet of S. putrefaciens formed
when media was spinned down. ............................................................................................................ 133
Figure S10-2 Representative cyclic voltammograms for Shewanella putrefaciens biofilms grown in the presence
of (non-basal, e.g. 0.1 μM) higher levels of Riboflavin (1 μM). Respective first Derivatives of each
voltammogram are also shown, scan rate 1 mV s-1
. .............................................................................. 134
-xii-
Figure S10-3 Effect of the Riboflavin concentration in the extracellular electron transfer. Representative cyclic
voltammogram of a Shewanella putrefaciens biofilm grown at a poised (+0.4 vs Ag/AgCl) graphite
electrode. The basal concentration of Riboflavin in the growth media was 0.1 μM as reported in the
Materials and Methods section (left panel). The voltammogram was recorded at maximum biofilm
activity after the start of the chronoamperometry with a scan rate of 1 mV s-1
. Voltammetry of all
Shewanella biofilms grown at different applied potentials with no additional supplementation of
Riboflavin (0.1 μM) showed only one inflection point centered at 0 V (vs Ag/AgCl). After six semi
batch chronoamperometric cycles a pulse of fresh substrate containing 1 μM of Riboflavin was injected
into the electrochemical cell (right panel). For the experiment with additional Riboflavin (1 μM) not
only the inflection point at 0 V was observed but also an inflection point centered at -0.4 V
characteristic of the mediator molecule Riboflavin (Peng, et al., 2010b), indicating that this molecule
participated in the extracellular electron transfer process. Furthermore, from the pronounced sharp rise
of the inflection point centered at the midpoint potential of Riboflavin, provided an example of how this
mediator molecule increases the electron transfer (Marsili, et al., 2008a). ........................................... 135
Figure S11-1 Influence of the buffer capacity: Cyclic voltammogramms (1mV s-1
) at pH 7, wastewater derived
and acetate–fed biofilms at varying buffer concentration, A) non-turn over B) turn over conditions. . 136
Figure S11-2 T-RFLP chromatograms (restriction digestion with RsaI and MspI) of the anode biofilms formed at
pH 7. The x axis represents the length of terminal restriction fragments and the y axis the relative
fluorescence units. On the right the area of every peak is shown as percentage of the total peak area. The
RsaI peak at 238 bp (503 bp with MspI) is shown in bright yellow and represents Geobacter
sulfurreducens (identified after sequencing). ........................................................................................ 137
Figure S11-3 T-RFLP chromatograms (restriction digestion with RsaI and MspI) of the anode biofilms formed at
pH 9. The x axis represents the length of terminal restriction fragments and the y axis the relative
fluorescence units. On the right the area of every peak is shown as percentage of the total peak area. The
peak at 238 bp (503 bp with MspI) is shown in bright yellow and represents Geobacter sulfurreducens
(identified after sequencing). In the sample of electrode-set 2 this organism could not be detected. This
biofilm comprised several phylotypes. ................................................................................................. 138
Figure S11-4 T-RFLP chromatograms (restriction digestion with RsaI and MspI) of the anode biofilms formed at
pH 6. The x axis represents the length of terminal restriction fragments and the y axis the relative
fluorescence units. On the right the area of every peak is shown as percentage of the total peak area. The
RsaI peak at 238 bp in the electrode-set 2 is shown in bright yellow and represents Geobacter
sulfurreducens (identified after sequencing the sample of electrode-set 2). In the small dashed window
the peak position is drawn to a larger scale to see that the peak position of the RsaI peak is different in
the sample of set 1 and set 2. The main MspI peak is found at 161 bp that is also different from what
was found for Geobacter sulfurreducens in the other samples (Figures S11-2 and S11-3 above). This
clearly shows that Geobacter sulfurreducens could not be detected in the sample of electrode-set 1. This
biofilm comprised several phylotypes. ................................................................................................. 139
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Figure S11-5 T-RFLP chromatograms (electrode-set 2, restriction digestion with RsaI) of the replenished
medium at the different feeding cycles. On the right the area of every peak is shown as percentage of
the total area. The peak at 238 bp is represented in bright yellow colour. It was only found in samples of
the feeding cycles at pH 7 and not in those at pH 9 (less than 1%). In this figure, in comparison to the
Fig. S11-2 above, a different resolution on the y axis was chosen to give a better overview of the present
diversity. Equal amounts of DNA were used for the analysis of all samples. ....................................... 140
Figure S11-6 Similarity analysis derived from anode chamber communities when treated over respective feeding
cycles at pH 7 and 9 (all electrode set 2). As can be observed, the T-RFLP derived composition of the
pH 7 and 9 communities was clearly different. Undoubtedly, the electrode biofilms were similar in T-
RFLP composition for pH 6 and 7 whereas the biofilm composition on the electrode treated at pH 9 was
different (Analysis: non-metric MDS, similarity measure: Bray-Curtis). ............................................. 141
Figure S11-7 Analysis of community structure by measuring the cells’ DNA contents and Forward scatter
behavior. Samples were harvested from the pH 9 anode chamber (electrode-set 2). ............................ 142
Figure S11-8 Analysis of community structure by measuring the cells’ DNA contents and Forward scatter
behavior. Samples were harvested from the pH 6 anode chamber (electrode set 2). ............................ 143
Figure S11-9 Cluster dendrogram derived from anode chamber communities when treated over several feeding
cycles and at different pH. Feeding cycle numbers and pH affiliation are given with c 1-5 and pH 6 to
pH 9 (shown for electrode-set 2). As can be observed, the structure of the inoculum community and that
of the pH 9 electrode are clearly different from all other samples. It is also obvious that distinct feeding
cycles cluster together such as pH 7 c1 to c3, pH 6 c2 to c4 and, pH 9 c2 to c4. It can be stated that
similar micro-environments like successive feeding cycles at a distinct pH value generated related
community structures. A few of the pH related communities clustered apart like pH 7 c4 to c5 or pH 6
c1 but are nevertheless close to each other if the similarity analysis of Figure S11-9 is included.
Undoubtedly, the electrode biofilms were similar in structure for pH 6 and pH 7. .............................. 144
Figure S11-10 Illustration of methodology used for estimating community similarities of cytometric flow plots
using a Dalmatian-plot. Areas of gates were estimated as sum of pixels for presence-absence when cell
abundances taken into account. Sums were calculated from plots of each of the samples separately and
for the overlap of two samples, respectively. For similarity estimation a modified Jaccard index was
used (Figure S11-10 taken from (Müller, et al., 2011). ......................................................................... 146
Figure S11-11 Photograph of the detachment of a pH 7 grown biofilm from an electrode due to extreme pH-
conditions (pH 11). ............................................................................................................................... 146
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Index of tables
Table 1-1 Representative microbially produced redox mediators. ........................................................................ 10
Table 1-2 Common terminology for the BES technology..................................................................................... 14
Table 2-1 Summary of the studied mutants and the achieved maximum current densities per projected electrode
surface area, the literature data are the reported maximum current densities in MFC experiments at
constant resistances. ................................................................................................................................ 38
Table 2-2 Result of the CV subtraction analysis (details in Fig. 5 and the text). .................................................. 45
Table 5-1 Cumulative data on electrocatalytic current densities obtained at different electrode materials. The
substrate was 10 mM Sodium acetate. .................................................................................................... 76
Table 6-1 Properties and anodic performance of carbon fiber mats. ..................................................................... 85
Table S9-1 Comparison of geometric current densities for Shewanella oneidensis Wild-type in different studies.
.............................................................................................................................................................. 120
Table S10-1 Comparison of geometric current densities for different strains of Shewanellaceae. ..................... 130
Table S10-2 Shewanella strains used as comparison in Table S10-1 and a description of their isolation
environment. ......................................................................................................................................... 132
Table S10-3 Cathodic and anodic peak positions, formal potential (vs. Ag/AgCl) and width of potential window,
ΔE, at a scan rate of 1 mV s-1
after SOAS baseline correction. ............................................................ 132
- 1 -
CHAPTER I
1 Extracellular electron transfer in Bioelectrochemical
systems: bridge between natural environments and
applied technologies
1.1 Prelude
In this introductory chapter a comprehensive description of microbial electron transfer
mechanisms in anoxic natural environments and the application of this natural process into a
promising, multi interdisciplinary -and still in continuing development technology- is given.
Section 1.2 illustrates the ecological significance of insoluble metal electron acceptors in nature.
Iron is taken as a model example to explain its bio-mobility in the environment. Here the
participation of some exemplary microorganisms capable of reducing iron is described. Section
1.3 provides a general definition of microbial extracellular electron transfer (ET) and describes
how microbiologists discovered this process in two model microorganisms now commonly used
as exemplary dissimilatory metal reducing bacteria. Later, one of the first applications for ET in
the field of bio-remediation and more recently in the field of Bioelectrochemical systems (BESs)
is provided. BESs not only have allowed the study of microbial ET but also permitted the
development of promising applications. Section 1.4 presents two known ET mechanisms
performed by bacteria, i.e., direct and mediated extracellular electron transfer (DET and MET
respectively). For DET, detailed descriptions on representative dissimilatory metal reducing
bacteria are given. In the case of MET, mediating redox species that transfer electrons between
the bacteria and the final electron acceptor are presented. Section 1.5 gives an overall
introduction to BESs. First, BESs represent an additional approach for the study of microbial ET
and second, they have emerged as an applied technology based on microbial ET. Finally Section
1.6 provides a comprehensive view on one of the main motivations in the development of BESs:
the improvement of current density production focused for near future applications. Different
aspects are exemplified with the case of 3D new electrode materials that improve the overall
performance of BESs. Finally, several environmental factors affecting the formation and
performance of electroactive biofilms are discussed.
-2-
1.2 Ecological significance of insoluble metal electron acceptors: the example of iron
Until the late 70s, reduction of Fe(III) to Fe(II) in sedimentary and subsurface environments was
believed to be the result of purely abiotic processes (Cornell and Schwertmann, 2007, Fenchel
and Blackburn, 1979). Now it is known that bacterial utilization of Fe(III) oxides as the terminal
electron acceptor is an important practice in anaerobic environments in which the reduction of
Fe(III) to Fe(II) is a enzymatically catalyzed bacterial process (Gralnick and Newman, 2007,
Lovley, 1993). Bacterial reduction of Fe(III) oxides has diverse significant ecological
repercussions, for example the quality of water can be modified by the increment of dissolved
Fe(II) that changes the taste of drinking water (Lovley, 2000) and furthermore Fe(III) is thought
to be the most abundant of all the available terminal electron acceptors in several subsurface
environments (Lovley, 1991). Some known representative microorganisms capable of utilizing
iron as final electron acceptor include: Geobacter metallireducens (Lovley, 1993),
Desulfuromonas acetoxidans (Roden and Lovley, 1993), Pelobacter carbinolicus (Lovley, et al.,
1995), members of the genus Desulfuromusa (Fredrickson and Gorby, 1996), Shewanella
oneidensis (Myers and Nealson, 1988), Ferrimonas balearica (Lovley, 2000), Geovibrio
ferrireducens (Caccavo Jr, et al., 1996) and Geothrix fermentans (Coates, et al., 1999).
The reduction of Fe(III) is considered as a predominant process due to the iron cycle reactions
(Lovley, et al., 1993), some of them with an important participation of bacteria (see below).
According to Luu and Ramsay (Luu and Ramsay, 2003), first solid oxides settle into the oxygen
transition zone called suboxic zone (Fig. 1-1). Simultaneously phosphate and metals are
removed via precipitation and complexation. In the suboxic zone carbon oxidation takes place
by bacteria via the utilization of iron as terminal electron acceptor. During iron reduction,
organic phosphate and metals are released into the oxic zone. From the oxidation of carbon,
Fe(II) forms insoluble precipitates in the suboxic zone such as siderite (FeCO3), pyrite (FeS2),
vivianite [Fe3(PO4)2] and magnetite (Fe3O4). Additionally some species of Fe(II) diffuse into the
oxic zone where finally reoxidation of Fe(II) occurs to form insoluble oxides and if no input of
organic carbon takes place, oxides accumulate in sediments of the suboxic zone, otherwise the
cycle continues again. Since the distribution of Fe(III) in the environment depends on the
amount of organic matter present (Pan, et al., 2011), Fe(III) oxides get retained in the sediment
when no organic matter is available diminishing the cycling of iron. Therefore the mobility of
certain compounds in the environment mainly depends on the biotransformation of organic
matter by microorganisms, making the study of these processes of great importance.
-3-
Figure 1-1 Simplified iron cycle in aquatic environments. Figure drawn with modifications after
(Luu and Ramsay, 2003, Nealson and Saffarini, 1994).
1.3 Electron transfer processes in the environment
Extracellular electron transfer (ET) is a general mechanism by which microorganisms generate
energy for cell growth and maintenance (Hernandez and Newman, 2001), i.e., bacteria transfer
electrons from their internal metabolism through a chain of trans-membrane proteins to finally
reduce insoluble metal electron acceptors. In the early 90s, environmental microbiologists
realized the importance of microbial ET to insoluble metal electron acceptors in several
biogeochemical cycles and progressively applied this extraordinary finding, e.g., on the
bioremediation of contaminated sites (Lovley, 1991, Nealson, et al., 1991). More recently this
finding has been used in an interdisciplinary way not only to study the fundamentals of
microbial ET but also to apply this concept in the so-called Bioelectrochemical systems (BESs)
(Rabaey, et al., 2010) (section 1.5). The basic and applied interest on microbial ET has rapidly
increased since the publication of two breakthrough papers introducing two of the first known
bacteria capable of reducing insoluble metal electron acceptors: Shewanella oneidensis MR-1
(Myers and Nealson, 1988) and Geobacter sulfurreducens PCA (Caccavo, et al., 1994).
-4-
Furthermore, the exploration of how microbes breathe minerals has been later stimulated by the
publication of both genomes (Heidelberg, et al., 2002, Methé, et al., 2003), making possible
genetic manipulations to study their respective ET pathways (see Chapter 2 for an example on
Shewanella oneidensis MR-1 knock-out mutants).
1.4 Microbial extracellular electron transfer mechanisms
To date mainly two microbial ET mechanisms have been recognized in the literature (Gralnick
and Newman, 2007, Hernandez and Newman, 2001, Lovley, 2011, Schröder, 2007, Watanabe,
et al., 2009). In one of those mechanisms named as direct extracellular electron transfer (DET),
electrons are transferred from the respiratory chain in the cell to an extracellular insoluble
compound or final electron acceptor (e.g., iron oxides or conductive electrode materials in
BESs) via a complex architecture involving several outer membrane cytochromes (Millo, et al.,
2011) (Fig 1-2A), an ability often conventionally awarded only to gram-negative bacteria
(Hernandez and Newman, 2001, Lovley, 2008a, Rosenbaum, et al., 2011, Shi, et al., 2009) with
some recent exceptions of gram-positive bacteria (Cournet, et al., 2010, Marshall and May,
2009, Wrighton, et al., 2011).
Figure 1-2 Overall illustration of microbial ET mechanisms found in the literature. A) Direct
extracellular electron transfer via membrane bound cytochromes and conductive nanowires and
B) Mediated extracellular electron transfer via a mediator molecule (Medred
or Medox
) (see text).
Here ET mechanisms are represented in the field of BESs with electrode materials as final
electron acceptors but the same illustration could be applied for bacteria in natural environments
using for instance iron oxides as terminal electron acceptors. Figure drawn with modifications
after (Schröder, 2007).
-5-
Another well-considered DET mechanism which is still under investigation is the ET via
cellular appendages facing the extracellular environment (i.e., microbial nanowires) found in
several bacteria (Bretschger, et al., 2010b) (Fig 1-2A) (see section 1.4.1). On the other side,
microorganisms are capable of ET via mediator molecules that, i) get reduced by outer
membrane cytochromes and later oxidized onto extracellular insoluble compounds or onto
conductive electrode materials as in the case of BESs; or ii) via periplasmatic or cytoplasmatic
redox couples that serve as reversible terminal electron acceptors, transferring electrons from the
bacterial cell to a final electron acceptor (Schröder, 2007). This mechanism is usually named as
mediated extracellular electron transfer (MET) (Marsili and Zhang, 2010) (Fig 1-2B) (see
section 1.4.2).
1.4.1 Microbial direct extracellular electron transfer (DET)
1.4.1.1 DET via membrane-bound redox-enzymes
As pointed out in section 1.2, diverse groups of microorganisms are now known to engage in
electron transfer to extracellular insoluble compounds. More recently with the use of conductive
electrode materials (anodes) in BESs, an additional number of microorganisms have joined to
the list of -recently named- exoelectrogenic bacteria capable of performing DET (Logan, 2009);
e.g., Desulfuromonas acetoxidans (Bond, et al., 2002), Escherichia coli K12 (Schröder, et al.,
2003), Rhodoferax ferrireducens (Chaudhuri and Lovley, 2003), Aeromonas hydrophila (Pham,
et al., 2003), Desulfobulbus propionicus (Holmes, et al., 2004a), Hansenula anomala (Prasad, et
al., 2007), Rhodopseudomonas palustris DX-1 (Xing, et al., 2008), Klebsiella pneumoniae L17
(Zhang, et al., 2008) and Proteus vulgaris (Rawson, et al., 2011) among others. While it is
commonly accepted that microbial ET occurs within complex communities found in BES
anodes (Logan and Regan, 2006a), the in-depth study of microbial ET mechanisms has revolved
around two model exoelectrogenic bacteria families: Shewanellaceae and Geobacteraceae
(Bretschger, et al., 2010b).
1.4.1.1.1 Shewanella oneidensis DET via membrane-bound redox-enzymes
As recently reported by Shi and co-workers (Shi, et al., 2009), DET performed by Shewanella
oneidensis depends on inner (IM) and outer membrane (OM) proteins that are known to be
directly involved in the reduction of insoluble metals that act as extracellular electron acceptors
(or in the case of BESs: electrode materials). These proteins include the inner membrane
tetrahaem c-Cyt CymA that is a homologue of NapC/NirT family of quinol dehydrogenases, the
-6-
periplasmic decahaem c-Cyt MtrA, the outer membrane protein MtrB and the OM decahaem c-
Cyts MtrC and OmcA (Fig. 1-3A).
All these proteins together form a pathway to transfer electrons from the quinone/quinol pool in
the inner membrane to the periplasm (PS) and then to the outer membrane where MtrC and
OmcA can transfer electrons directly to the surface of electrode materials.
1.4.1.1.2 Geobacter sulfurreducens DET via membrane-bound redox-enzymes
On the other side, DET performed by Geobacter sulfurreducens (as reported by Shi and co-
workers (Shi, et al., 2009)) relies on the outer membrane proteins tetrahaem c-Cyt OmcE and
hexahaem c-Cyt OmcS that are believed to be located on the cell surface where they are
suggested to transfer electrons to type IV pili. Type IV pili are hypothesized to transfer electrons
directly to Fe(III) oxides (or in the case of BESs: electrode materials). OmcE and OmcS also
receive the electrons from the quinone/quinol pool in the inner membrane (Fig. 1-3B).
Figure 1-3 Roles of outer membrane cytochromes of A) Shewanella oneidensis and B)
Geobacter sulfurreducens in extracellular electron transfer. IM: inner membrane, OM: outer
membrane and PS: periplasm. Figure drawn with modifications after (Shi, et al., 2009).
1.4.1.2 DET via self-produced microbial nanowires
The fundamental comprehension of microbial ET mechanisms is still in progress (Bretschger, et
al., 2010b) since non-conclusive and debatable experimental evidence of an additional DET
process via self-produced microbial nanowires has come to light (Lovley, 2011). This recently
found DET mechanism is not only expected to change the way scientists will look at microbial-
-7-
electrode interactions but also it could commence a new whole applied research field due to the
promising application of microbial nanowires as nano bio-conductive materials (Malvankar, et
al., 2011). In general, the information devoted to the analysis of conductive bacterial nanowires
is scarce. However experimental evidence of microbial-like nanowires has been reported for
some microorganisms as described below. There exists evidence showing the presence of
microbial-like nanowires in nutrient limited cultures of the cyanobacterium Synechocystis sp.
PCC 6803 (Fig. 1-4C) and in co-cultures of Pelotomaculum thermopropionicum and
Methanothermobacter thermautotrophicus (Fig. 1-4D) (Gorby, et al., 2006). Additionally,
putative nanowires have been observed in sulfate limiting cultures of Desulfovibrio vulgaris and
in environmental samples from hydrothermal vents. Nevertheless, only visual information in
this regard has been presented so far (Bretschger, et al., 2010b). Whereas microbial-like
nanowires structures have been observed in several bacterial cultures (Bretschger, et al., 2010b),
hitherto; to the best of my knowledge and beyond the optical description, only four works
devoted to the electrochemical and spectroscopical characterization of these structures have
been published (according to “Scopus”, February 2012) and all of them using either the model
exoelectrogenic bacterium G. sulfurreducens or S. oneidensis.
1.4.1.2.1 Geobacter sulfurreducens DET via self-produced microbial nanowires
One of the first observations on microbial nanowires was made by Reguera and co-workers
(Reguera, et al., 2005) on G. sulfurreducens. They have found that a nanowire-deficient mutant
of G. sulfurreducens could not reduce Fe(III). Additionally by using atomic force microscopy
they suggested that these G. sulfurreducens nanowires could be conductive. A few years later,
additional information on the possible conductivity of G. sulfurreducens nanowires was
provided by Malvankar and co-workers (Malvankar, et al., 2011). They have showed the
metallic-like conductivity (along centimeter-length scale) in microbial nanowires produced by
G. sulfurreducens. Moreover, they have even suggested that these structures could possess
similar properties to those of synthetic metallic nanostructures (Fig. 1-4A).
1.4.1.2.2 Shewanella oneidensis DET via self-produced microbial nanowires
On the other hand, only one year later to the first finding of nanowires in G. sulfurreducens,
Gorby and co-workers provided evidence on the conductivity of electrical microbial nanowires
produced by S. oneidensis in direct response to electron-acceptor limitations (Gorby, et al.,
2006). Four years later El-Naggar and co-workers (El-Naggar, et al., 2010) presented an
additional contribution in this regard confirming the conductivity of such microbial nanowires
produced by S. oneidensis MR-1 (Fig. 1-4B). Independent of the source of microbial nanowires,
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the experiments reported so far present the bacterial nanowires as a viable microbial strategy for
DET and more importantly represent a promising alternative for future nano bio-conductive
materials. Ultimately, although DET (via membrane-bound redox-enzymes or via microbial
nanowires) seems to be an imperative microbial ET mechanism in some species of
microorganisms, mediated electron transfer (MET, explained in the following section) via
mediator molecules has been proved as well to have an outstanding participation in the overall
ET process (see Chapter 2).
Figure 1-4 Scanning electron microscope micrographs of: A) Geobacter sulfurreducens (ATCC
51573) (Malvankar, et al., 2011); B) Shewanella oneidensis MR-1 (Gorby, et al., 2006); C)
Synechocystis sp. PCC 6803 (Gorby, et al., 2006) and D) co-culture of Pelotomaculum
thermopropionicum and Methanothermobacter thermautotrophicus showing nanowires
connecting the two genera (Gorby, et al., 2006).
1.4.2 Microbial mediated extracellular electron transfer (MET)
Microbial mediated extracellular electron transfer (MET) requires transfer of electrons from the
respiratory chain in the cell to extracellular inorganic material via a redox mediator molecule.
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The known microbial MET occur via i) artificial exogenous mediator molecules; ii) natural
exogenous mediator molecules; and iii) self-produced mediator molecules.
1.4.2.1 MET via artificial exogenous mediator molecules
In early experiments with BESs, the need of exogenous mediator molecules was believed to be
crucial for bacteria to transfer electrons to electrodes immersed in bacterial solutions (Cohen,
1931). The approach of using these molecules was applied again in the 1980s mainly by
Bennetto and co-workers (Bennetto, et al., 1983). The majority of mediator molecules were
based on phenazines (Park and Zeikus, 2000), phenothiazines (Delaney, et al., 1984),
phenoxazines (Bennetto, et al., 1983) and quinones (Tanaka, et al., 1988) demonstrating their
suitability as redox mediators between certain bacteria and electrode materials. More recently,
additional compounds have been reported as well, e.g.: resazurin (Sund, et al., 2007), humate
analog anthraquinone 2-6-disulfonate (Milliken and May, 2007) and methyl viologen (Aulenta,
et al., 2007). Although exogenous mediator molecules are easy to dose and their redox potential
may be adjusted over a wide range by careful design of the molecule (Marsili and Zhang, 2010),
their main disadvantage is the necessity of a regular addition of these compounds, which from a
practical point of view is technologically unfeasible and environmentally questionable
(Schröder, 2007).
1.4.2.2 MET via natural exogenous mediator molecules
In MET, microbes can use natural exogenous (non self-produced) electron shuttling compounds
available in the subsurface environment such as humic acids (Fredrickson, et al., 2000a,
Fredrickson, et al., 2000b, Lovley, et al., 1996, Straub, et al., 2005), cysteine (Doong and
Schink, 2002, Kaden, et al., 2002) or sulfur-containing compounds (Straub and Schink, 2003).
The importance of such natural exogenous mediator molecules lies in the fact that this kind of
molecules have found to be responsible for MET in natural sediments (Nielsen, et al., 2010).
1.4.2.3 MET via self-produced mediator molecules
Finally and more importantly (from the ecological and applied point of view), it is assumed that
microorganisms due to environmental restriction use endogenous redox mediators (self-
produced by bacteria) to accomplish the production of energy for cell growth and maintenance
by the reduction of insoluble terminal electron acceptors. Initial experiments to produce and
characterize mediator molecules were done through insoluble metal reduction assays (Caccavo,
et al., 1994, Myers and Nealson, 1988). Only relatively recently, the use of BESs (see Section
-10-
1.5) has stimulated the general interest on externally microbial ET (Bond, et al., 2002, Kim, et
al., 1999a).
To date, mainly experiments with gram-negative bacteria have contributed with evidence that
microorganisms are able to perform MET mechanisms (Marsili and Zhang, 2010). Microbial
known mediators are listed in Table 1-1. In general, these molecules have provided experimental
evidence on the possibility to transfer electrons to electrode materials and according to
assumptions made by Marsili and Zhang (Marsili and Zhang, 2010), redox mediator molecules
would be able to transfer electrons between bacteria and final electron acceptors regardless of a
solid metal oxide or an electrode material. Such an ability in conjunction with the fact that self-
produced mediator molecules from one bacteria can be used further by a different bacteria (as in
the case of Pseudomonas sp. and Brevibacillus sp. PTH1 (Pham, et al., 2008)) increases the
applications of this specific MET mechanism.
Table 1-1 Representative microbially produced redox mediators.
Microoganism Mediator molecule Reference
Sphingomonas xenophaga 4-amino-1,2-naphthoquinone
(Keck, et al., 2002)
Pseudomonas aeruginosa Phenazine-1-carboxylic acid
(Price-Whelan, et al., 2006)
Pseudomonas chlororaphis Phenazine-1-carboxamide
(van Rij, et al., 2004)
Shewanella oneidensis Flavin mononucleotide
(von Canstein, et al., 2008)
Shewanella algae Melanin
(Turick, et al., 2002)
Bacillus pyocyaneus Pyocyanine
(Friedheim and Michaelis,
1931)
Propionibacterium
freundenreichii
2-Amino-3-carboxy-1,4-
naphthoquinone
(Hernandez and Newman,
2001)
Shewanella alga Cyanocobalamin
(Workman, et al., 1997)
Acetobacterium woodii Hydroxycobalamin (Hashsham and Freedman,
1999)
Pseudomonas stutzeri Pyridine-2,6-bis
(Lewis, et al., 2001)
Methanosarcina thermophila Porphorinogen-type molecules
(Koons, et al., 2001)
Geobacter metallireducens Anthraquinone-2,6-disulfonate
(Cervantes, et al., 2004)
Shewanella oneidensis 1,4-Dihydroxy-2-naphthoate
derivative (Ward, et al., 2004)
Klebsiella pneumoniae Anthraquinone-2,6-disulfonate
(Li, et al., 2009b)
aMore detailed information can be found in the following references: (Hernandez and Newman,
2001, Marsili and Zhang, 2010, Schröder, 2007, Watanabe, et al., 2009).
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1.5 Bioelectrochemical systems (BESs)
From Section 1.1 there has been a constant reference on BESs since these systems have
represented a driving force in the elucidation of microbial electron transfer mechanisms.
Although it could be assumed that microbial BESs represent a novel research field, this is not
completely true. The technology in fact is quite old and just recently has been revisited
(Schröder, 2011). The ability of microorganisms to transfer electrons from the internal
metabolic chains to extracellular terminal acceptors (with the concomitant production of an
electric current) was discovered more than 100 years ago (Schröder, 2011). However, this
finding has attracted increasing attention only during the last decade (Hernandez and Newman,
2001, Schroder, 2007, Watanabe, et al., 2009). Michael C. Potter reported in the year 1911 the
electromotoric force between electrodes immersed in bacterial cultures in a battery (Potter,
1911). In Potter’s communication, he concluded that electric energy could be generated from the
microbial decomposition of organic compounds. With this unusual (at that time) combination of
microbiology and electrochemistry, Potter was a pioneer providing one clearer hint on the
consequences of the bacterial metabolism. As reviewed in previous sections, microbial ET has
received great attention not only for the basic knowledge of how electrons end at an electron
acceptor from the geochemistry point of view but also for the possible use of this extraordinary
process in bioremediation, in the production of bioenergy and/ or more recently in the
production of valuable products by the so called BESs (Rabaey, et al., 2009, Rabaey and
Rozendal, 2010). Additionally, this interest has been clearly reflected by the number of
publications including the use of BESs (Fig. 1-5).
In BESs, a plenitude of possible applications can be found (Fig. 1-6), from the original and
promising production of electricity (Logan, et al., 2006), to hydrogen as a clean fuel (Logan, et
al., 2008) and the production of useful chemicals (Rabaey and Rozendal, 2010) such as
hydrogen peroxide, extraordinarily from wastewater (Fu, et al., 2010, You , et al., 2010).
Nonetheless, the cited applications in this section would not be possible without the basic
research on the microbe-electrode interactions which inexorably turn out to contribute to the
betterment of the overall performance of this kind of systems by eliminating (or at least
diminishing) electrochemical losses of BESs (Schröder and Harnisch, 2010). Therefore, the
analysis of the microbe-electrode interactions would lead not only to a higher comprehension on
improving the overall performance of BESs (see section 1.5) from the power production point of
view but also on improving a more precise electron uptake by microorganisms for the
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production of useful and industrial demanded biochemicals (Nevin, et al., 2010, Ross, et al.,
2011).
Figure 1-5 Number of publications reporting the use of “Bioelectrochemical systems” (Scopus
data base, January 2012). Illustration based on (Schröder, 2011).
As shown in Fig. 1-6, microbial-electrode interactions can take place in both electrode chambers
depending on the application for which the BES has been designed. A simplified version of a
BES system as shown in the insert of Fig. 1-6 is a potentiostatic controlled electrochemical half-
cell in which an anode and a cathode are hosted within one vessel (LaBelle, et al., 2010). This
experimental approach assures similar biological and environmental conditions for both
electrodes and increases the reproducibility of the experiment by maintaining one of the
electrodes at a constant potential permanently controlled against a reference electrode (e.g., vs.
Ag/AgCl) (Bard, et al., 2008). This type of BES (with multiple modifications) is the one that has
been extensively used in this Thesis.
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Figure 1-6 Overall view of Bioelectrochemical systems. Production of electricity and useful
metabolites take place in BESs. These microbial/ enzyme/ organelles based systems consist of
an anode (oxidation process), a cathode (reduction process) and typically a membrane separating
both electrodes (see also Table 1-2). Depending on the membrane specificity (Harnisch and
Schröder, 2009), type of catalysts at both electrodes (Franks, et al., 2010, Rosenbaum, et al.,
2011), and the source of the reducing power (Logan, et al., 2008, Logan, et al., 2006) a diverse
spectrum of research and practical applications can be found (see Section 1.5.1). Drawn with
modifications after (Rabaey and Rozendal, 2010).
1.5.1 Types of Bioelectrochemical systems
Depending on the application, the BES receives a different name s as seen in Table 1-2. From
the different BESs that can be found in the literature, only a few of them have attracted most of
the scientific community’s attention, e.g.: microbial fuel cells (MFCs), microbial electrolysis
cells (MECs), microbial desalination cells (MDCs), microbial solar cells (MSC) and enzymatic
fuel cells (EFCs).
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Table 1-2 Common terminology for the BES technology.
Name Abbrev. Definition Ref.*
Bioelectrochemical
system
BES An electrochemical system in which biocatalysts
(microorganisms) perform oxidation and/ or reduction at
electrodes
[1]
Microbial fuel cell
MFC A BES that produces net electrical power [2]
Microbial electrolysis
cell
MEC A BES to which net electrical power is provided to achieve
a certain process or product formation
[3]
Bioelectrochemically
assisted microbial
reactor
BEAMR A BES to which net electrical power is provided to achieve
a certain process or product formation
[4]
Bio-electrical reactor
BER A reactor in which current is provided to microorganisms
to stimulate their metabolism
[5]
Biocatalyzed
electrolysis cell
BEC A BES to which net electrical power is provided to achieve
a certain process or product formation
[6]
Biochemical fuel cell
BFC An electrochemical system in which biocatalysts function
as catalysts for oxidation and/ or reduction reaction at
electrodes
[7]
Biofuel cell
BFC An electrochemical system that use biocatalysts to convert
chemical energy to electrical energy
[8]
Sediment microbial
fuel cell
SMFC MFC operated at underwater sediment interface [9]
Benthic unattended
generator
BUG MFC operated at underwater sediment interface [10]
Enzymatic fuel cell
EFC An electrochemical system in which biocatalysts
(enzymes) perform oxidation and/ or reduction at
electrodes
[11]
Microbial desalination
cell
MDC An MFC for desalinating water based on using the
electrical current generated by exoelectrogenic bacteria
[12]
Microbial solar cell
MSC An MFC that exploits the energy of light and the activity
of phototrophic microorganisms to produce electricity
[13]
Mitochondrial biofuel
cell
MBFC A new class of BES that uses whole organelles (e.g.,
mitochondria) as catalysts
[14]
Note: Table based on information available in (Rabaey, et al., 2010). *References in Table: 1: (Rabaey,
et al., 2007); 2: (Logan, et al., 2006); 3: (Logan, et al., 2008); 4: (Ditzig, et al., 2007); 5: (Thrash and
Coates, 2008); 6: (Rozendal, et al., 2006b); 7: (Lewis, 1966); 8: (Cooney, et al., 2008); 9: (Reimers, et
al., 2000); 10: (Lovley, 2006); 11: (Minteer, et al., 2007); 12: (Kim and Logan, 2011); 13: (Rosenbaum
and Schröder, 2010); 14: (Bhatnagar, et al., 2011).
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1.5.1.1 Microbial fuel cells
As a general definition, microbial fuel cells (MFCs) are devices that use bacteria as the catalysts
to oxidize organic and inorganic matter and generate current (Logan, et al., 2006). According to
Logan and co-workers (Logan, et al., 2006), in a MFC bacteria oxidize organic matter and
release carbon dioxide and protons into solution and electrons to an anode. Electrons are then
transferred by DET or MET to the anode (or working electrode) and flow to the cathode (or
counter electrode) linked by a conductive material containing a resistor, or operated under a load
(see Fig. 1-6). Finally, the electrons that are transferred from the anode to the cathode combine
with protons (that diffuse from the anode chamber through a physical separator) and oxygen
provided from air to produce water.
1.5.1.2 Microbial electrolysis cells
Unlike MFCs, Microbial electrolysis cells (MECs) use electrochemically active bacteria to
break down organic matter, combined with the addition of a small voltage that results in
production of hydrogen gas (Logan, et al., 2008). MECs used to produce hydrogen gas are
similar in design to MFCs that produce power, but there are important differences. According to
Logan and co-workers (Logan, et al., 2008) in a MFC, when oxygen is present at the cathode,
current can be produced, but without oxygen, current generation is not spontaneous. However, if
a small voltage is applied, current generation is forced between both electrodes and hydrogen
gas is produced at the cathode through the reduction of protons.
1.5.1.3 Microbial desalination cells
Microbial desalination cells (MDCs) are based on transfer of ionic species out of water in
proportion to current generated by bacteria (Luo, et al., 2012). Developed by Cao and co-
workers (Cao, et al., 2009), MDCs consist of three chambers, with an anion exchange membrane
next to the anode and a cation exchange membrane by the cathode, and a middle chamber
between the membranes filled with water that is being desalinated. When current is generated by
bacteria on the anode, and protons are released into solution, positively charged species are
prevented from leaving the anode by the anion exchange membrane and therefore negatively
charged species move from the middle chamber to the anode. In the cathode chamber protons
are consumed, resulting in positively charged species moving from the middle chamber to the
cathode chamber. This loss of ionic species from the middle chamber results in water
desalination.
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1.5.1.4 Microbial solar cells
When sunlight is converted into electricity within the metabolic reaction scheme of a MFC, this
system is described as photosynthetic MFC or microbial solar cell (MSC) (Rosenbaum, et al.,
2010b). MSCs are used to convert light into electricity by exploiting the photosynthetic activity
of living, phototrophic microorganisms (Rosenbaum and Schröder, 2010). These BESs have
been described in detail by Rosenbaum and co-workers (Rosenbaum, et al., 2010b). In their
publication they indentify five different approaches that integrate photosynthesis with MFCs: a)
photosynthetic bacteria at the anode with artificial mediating redox species, b) hydrogen-
generating photosynthetic bacteria with an electrocatalytic anode, c) photosynthesis coupled
with mixed heterotrophic bacteria at the anode, d) direct electron transfer between
photosynthetic bacteria and electrodes and e) photosynthesis at the cathode to provide oxygen.
1.5.1.5 Enzymatic fuel cells
Enzymatic fuel cells (EFCs) are energy conversion devices that use enzymes as biocatalysts to
convert chemical energy to electrical energy (Cooney, et al., 2008). According to Cooney and
co-workers (Cooney, et al., 2008), BESs are usually classified on the basis of the type of
biocatalyst employed. There are three types of biocatalyst used in BESs: microbes, organelles,
and enzymes, each of this type has advantages and disadvantages. While MFCs can operate for
years (Logan, 2010) and completely oxidize their fuel, MFCs have been limited by low current
and power densities. On the other hand, EFCs have been shown to have higher current and
power densities, but have been limited by incomplete oxidation of fuel and lower active lifetime
(Minteer, et al., 2007).
1.6 Performance of Bioelectrochemical systems
As one can see from the literature (Schröder, 2011), one of the motivations for the development
of the BES technology has been a competitive “race” to increase the current production and
trying to make this technology an affordable option for the treatment of wastewater with the
concomitant consequence production of sustainable electricity and biochemicals (Rabaey and
Rozendal, 2010).
Here, the understanding of microbial-electrode interactions has been part of the global effort to
accomplish BESs with an enhanced performance. Current density based on available anode
surface area has made a noticeable development (Fig 1-7). Since 1999, the experimental
biotransformation of substrate (fuel) to electric energy (Schröder, 2007) has been performed
with the utilization of dissimilatory metal reducing bacteria (e.g., from the Shewanellaceae
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family (Kim, et al., 1999b, Kim, et al., 1999d)). The performance of the current density
production has seen a considerable increment from only 0.013 μA cm-2
(Kim, et al., 1999d) to
more than 30 A m-2
(see Chapter 5 and 6).
Figure 1-7 Illustration of the enhancement of the anodic current density performance in BESs.
Current density values taken from representative literature data: (Aelterman, et al., 2006, Bond,
et al., 2002, Catal, et al., 2008a, Catal, et al., 2008b, Chen, et al., 2011, Gil, et al., 2003, He, et
al., 2011, He, et al., 2005, Holmes, et al., 2004b, Katuri, et al., 2010, Kim, et al., 1999b, Kim, et
al., 1999d, Liu, et al., 2005, Liu, et al., 2010c, Milliken and May, 2007, Min and Logan, 2004,
Park and Zeikus, 2000, Park, et al., 2001, Torres, et al., 2009, Zhao, et al., 2010b, Zuo, et al.,
2006). Illustration based on Ref. (Schröder, 2011).
The betterment of performance of BESs based on the current density is (among other factors)
due to: i. the fabrication of porous three dimensional materials that allow bacteria to take
advantage of higher electrode surface areas to release electrons (Katuri, et al., 2011,
Šefčovičová, et al., 2011, Xie, et al., 2011, Yu, et al., 2011) (see Chapter 5 and 6); ii. the
comprehension of how electrochemically active bacteria associate with some electrode materials
through improved anode enrichment processes (Kim, et al., 2004, Liu, et al., 2008, Rabaey, et
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al., 2004); and iii. through the study of the process of biofilm formation influenced by
environmental factors (see Chapter 7 and 8).
1.6.1 Performance based on the improvement of electrode materials
Current density production in BESs has been always one of the most attractive objectives to be
achieved with these type of systems (Schröder, 2011) and as one can see from Fig. 1-7, the race
for improving the performance and finally making BESs an -on field- applied technology will
still continue (Keller, et al., 2010). To achieve this, contributions of the design of new materials
will be invaluable since these materials will have the challenge to enhance the microbe-electrode
interaction either by increasing the surface of contact between electroactive biofilms and
electrode materials or by allowing new electrode materials to collect more electrons effectively
from the internal metabolism of bacteria.
To date many strategies have been used in order to enhance the performance of BESs. These
strategies could be summarized as below:
i. improvement in the architecture design of BESs (Cheng, et al., 2006);
ii. increment of the buffer capacity in cathodic and anodic chambers (Fan, et al., 2008);
iii. use of respiratory inhibitors (Chang, et al., 2005);
iv. improved enrichment and acclimatization procedures of electroactive microbial biofilms
(Liu, et al., 2008);
v. construction of conductive artificial biofilms by the immobilization of electroactive bacteria
(Yu, et al., 2011); and just recently
vi. use of carbon based three dimensional electrode materials (Katuri, et al., 2011, Logan, et al.,
2007, Šefčovičová, et al., 2011, Xie, et al., 2011, Zhao, et al., 2010b).
In fact, commercially available carbon based materials are considered to be the most widely
used materials for BESs anodes due to their biocompatibility, chemical stability, high
conductivity, and relatively low cost (Wei, et al., 2011). All of these advantages have been
exploited in some recent reports that have succeeded in modifying these materials to enhance
the production of anodic current density (see below some examples).
For instance, Zhao and co-workers (Zhao, et al., 2010b) used a conductive polyaniline nanowire
network with three-dimensional nanosized porous structures as BESs anodes. They reported
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substantial improvements (10 to 100 fold) in current and power densities in comparison to
conventional two-dimensional materials (Schröder, 2011). More recently Katuri and co-workers
(Katuri, et al., 2011) fabricated three-dimensional microchannelled nanocomposite electrodes.
These materials allowed the growth of Geobacter sulfurreducens biofilms over the three-
dimensional surface, providing acetate oxidation current densities of up to 25 A m−2
. Xie and
co-workers (Xie, et al., 2011) reported a carbon nanotube sponge composite that provided a
three-dimensional scaffold that was favorable for microbial colonization. This nanotube sponge
allowed the increment of 2.5 times the previously reported maximum areal power density and 12
times the previously reported maximum volumetric power density.
Independently from the previous examples on carbon based three dimensional electrode
materials, Chapter 5 and 6 present two studies in this regard showing conditions that allowed the
production of the highest current density values reported so far by bio-electrochemically active
biofilms.
1.6.2 Performance based on the study of environmental factors affecting biofilm
formation
In the field of BESs, it has been assumed that the treatment of wastewater could be one of the
most appealing applications (Logan, et al., 2006). In fact, in order to make BESs a successful
technology in wastewater treatment, researchers have to pay special attention to the
environmental and external factors that influence the biofilm, considered to be “powerhouse” of
BESs (Franks, et al., 2010). In the literature one can find different approaches that have been
utilized in order to decipher the factors influencing the formation of anodic biofilms in BESs.
For instance, Patil and co-workers (Patil, et al., 2010) investigated the temperature dependence
and temperature limits of wastewater derived anodic microbial biofilms. They demonstrated that
these biofilms are active in a temperature range between 5 and 45°C. Additionally, they also
demonstrated that elevated temperatures during initial biofilm growth not only accelerated the
biofilm formation process but they also influenced the bioelectrocatalytic performance of these
biofilms when measured at identical operation temperatures. For example, the time required for
biofilm formation decreased from above 40 days at 15°C to 3.5 days at 35°C. On the other side,
Zhang and co-workers (Zhang, et al., 2011) investigated the effects of external resistance on
biofilm formation and electricity generation of microbial fuel cells. The morphology and
structure of the biofilms developed at 10, 50, 250 and 1000 Ω was characterized. They
demonstrated that the biofilm structure played a crucial role in the maximum power density and
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sustainable current generation of BESs. Their results showed that, maximum power density of
their BESs increased when the external resistance decreased. They have attributed their results
to the existence of void spaces beneficial for proton and buffer transport within the anode
biofilm, which maintains a suitable microenvironment for electrochemically active
microorganisms. Furthermore, Biffinger and co-workers (Biffinger, et al., 2009) used a high-
throughput voltage based screening assay to correlate current output from a BESs containing
Shewanella oneidensis MR-1 to biofilm coverage over 250 h (among other experimental
conditions). BESs operated by Biffinger and co-workers permitted data collection from nine
simultaneous S. oneidensis MR-1 BESs experiments in which each experiment was able to
demonstrate organic carbon source utilization and oxygen dependent biofilm formation on a
carbon electrode. Finally, Ieropoulos and co-workers (Ieropoulos, et al., 2010) have
hypothesized that the processing of large volumes of wastewater in BESs would require
anodophilic bacteria operating at high flow-rates. Therefore, they examined the effect of flow-
rate on different microbial consortia during anodic biofilm development using inocula designed
to enrich either aerobes/ facultative species anaerobes. By using scanning electron microscopy
they showed some variation in biofilm formation where clumpy growth was associated with
lower power. In a different category, experiments using genetic manipulations should be
mentioned. For example, the use of knocked mutants of bacteria in order to delete from their
genome the production of outer membrane surface structures needed to adhere to solid surfaces
and generate ticker and robust electroactive biofilms (Bouhenni, et al., 2010, Rollefson, et al.,
2009). Regardless of the previous examples on factors influencing the formation of anodic
biofilms in BESs, Chapter 7 and 8 present two more detailed examples on this aspect.
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1.7 Aim of this Dissertation
Because of the issues raised in the previous sections in this chapter, the aim of my dissertation
was to investigate different aspects of microbial-electrode interactions in BESs. The different
objectives of this Ph.D. Thesis are divided into the following chapters:
Part I Electron transfer mechanisms of pure culture biofilms of Shewanella spp.
Chapter 2 Cyclic voltammetric analysis of the electron transfer of Shewanella
oneidensis MR-1 and nanofilament and cytochrome knock-out mutants.
Chapter 3 Study of Shewanella putrefaciens biofilms grown at different applied
potentials using cyclic voltammetry and confocal laser scanning microscopy.
Chapter 4 Spectroelectrochemical analysis of intact microbial biofilms of Shewanella
putrefaciens for sustainable energy production.
Part II Porous 3D carbon as anode materials for performance of electrochemically active mixed
culture biofilms.
Chapter 5 Electrospun and solution blown three-dimensional carbon fiber nonwovens
for application as electrodes in microbial fuel cells.
Chapter 6 Electrospun carbon fiber mat with layered architecture for anode in microbial
fuel cells.
Part III The influence of external factors on electrochemically active mixed culture biofilms.
Chapter 7 Electroactive mixed culture biofilms in microbial bioelectrochemical
systems: the role of pH on biofilm formation, performance and composition.
Chapter 8 Electroactive mixed culture biofilms in microbial bioelectrochemical
systems: the role of the inoculum and substrate on biofilm formation, performance and
composition.
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1.8 Structure of the Thesis and personal contribution
This Ph.D. Thesis tackled various aspects within bioelectrochemical systems. For this reason,
several available experimental techniques were utilized. From electrochemical voltammetric
techniques, via confocal laser scanning microscopy, to surface-enhanced resonance Raman
scattering. The results of this Thesis are divided into three main parts regarding the respective
area of study:
Part I Electron transfer mechanisms of pure culture biofilms of Shewanella spp.
Part II Porous 3D carbon as anode materials for performance of electrochemically active mixed
culture biofilms.
Part III The influence of external factors on electrochemically active mixed culture biofilms.
From Fig. 1-8 one can see that the different parts of this Thesis were focused mainly on the
investigation of several processes occurring at the interface between the electrode material and
bioelectroactive biofilms. In the following lines, the chapters contained in this thesis are listed.
Furthermore an appreciation of my personal contribution to each is provided.
Figure 1-8 Schematic illustration of the research areas within the three chapter I, II and III.
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Part I. Electron transfer mechanisms of pure culture
biofilms of Shewanella spp.
CHAPTER 2
The idea of this project came as a continuation of previous experiments performed on
Shewanella at the University of Greifswald. The reason to use Shewanella oneidensis MR-1 as a
biological model came from a collaboration impelled by Dr. B.R. Ringeisen’s visit to our
Institute in November 2008. At that time Prof. Dr. U. Schröder and Dr. F. Harnisch motivated
me to work on electrochemical active biofilms of wild-type and mutant strains of S. oneidensis
kindly provided by Dr. B.R. Ringeisen’s team at the US Naval Research Laboratory. The
growth of Shewanella strains was performed in close collaboration with L.A. Fitzgerald and J.C.
Biffinger. I designed, planned and performed all experimental work in Braunschweig in close
collaboration with Dr. F. Harnisch. I analyzed/ interpreted data and wrote the manuscript
together with Dr. F. Harnisch. During the whole process of this project we maintained useful
discussions with Dr. B.R. Ringeisen’s team. Prof. Dr. U. Schröder gave useful advice.
CHAPTER 3
The lack of information on the electron transfer mechanisms of electrochemical active biofilms
of Shewanella initiated this study. Most of the previous studies were generally carried out with a
single applied potential or each study used different operational parameters, which makes it
difficult to compare among studies. I took care of the whole maintenance process and growth of
S. putrefaciens. I designed, planned and performed all experimental work in Braunschweig in
close collaboration with Dr. F. Harnisch and Prof. Dr. U. Schröder. CLSM measurements were
performed at the Helmholtz Centre, Magdeburg by U. Kuhlicke and me in close collaboration
with Dr. T.R. Neu. I analyzed/ interpreted data and wrote the manuscript together with Dr. F.
Harnisch. For the CLSM data we maintained invaluable discussions with Dr. T.R. Neu. Prof.
Dr. U. Schröder gave useful advice and guidance.
-24-
CHAPTER 4
The idea of this project came as a continuation of previous spectroelectrochemical experiments
performed in our group on Geobacter biofilms. The idea to use Shewanella putrefaciens as a
biological model came from the total lack of information on the electron transfer mechanisms
using spectroelectrochemical tools such as surface-enhanced resonance Raman scattering. This
project was conducted in several phases at the TU Braunschweig and at the TU Berlin. At the
TU Braunschweig, I took care of the whole maintenance process and growth of S. putrefaciens
biofilms in electrochemical half-cells. In close collaboration with Dr. D. Millo, I designed,
planned and performed experimental work at the TU Berlin in the group of Prof. Dr. P.
Hildebrandt. Under the supervision of Dr. Millo (who performed the SERRS experiments and
analyzed the spectra with the assistance of Khoa H. Ly), and Dr. Harnisch, I analyzed and
interpreted data. During the whole process of this project, Prof. Dr. U. Schröder has given useful
advice and guidance.
Part II. Porous 3D carbon as anode materials for
performance of electrochemically active mixed culture
biofilms
CHAPTER 5
The motivation of the study can be assigned to the continuous efforts in the field of BESs to
improve the overall performance of the systems; especially in terms of electrode materials, as
the bioelectrocatalytic activity plays a key role. The motivation for this project came from S.
Chen who was finishing at the time his Ph.D. at the Philipps-Universität in Marburg in the group
of Prof. Greiner. This project was conducted in several phases at the Universty of Marburg and
the TU Braunschweig. At the TU Braunschweig we tested a series of 3D porous carbon fiber
based materialss, produced by gas-assisted electrospinning at the Philipps-Universität and a
series of electrospun and solution-blown carbon fibers fabricated by the group of Prof. Yarin at
the University Illinois at Chicago on the suitability to serve as electrode materials for BESs. At
all times I was deeply involved in the growth and maintenance of waste-water derived
electroactive biofilms and the test of the mentioned electrode materials as well as data analysis
and interpretation.
-25-
CHAPTER 6
Encouraged by our previous work presented in detail in Chapter 5, this study was driven by the
continuous efforts in the field of BESs to develop high performance three-dimensional electrode
materials for electroactive biofilms. This project was conducted at the Philipps-Universität
Marburg in the group of Prof. Greiner (material development) and at the TU Braunschweig in
the group of Prof. Schröder (electrode characterization). We tested a series of electrospun
carbon fiber mats with layered architecture and investigated these materials on their suitability
for growth and performance of electroactive waste water derived anodic biofilms. At all times I
was deeply involved in the growth and maintenance of the waste-water derived electroactive
biofilms and the test of the mentioned electrode materials as well as data analysis and
interpretation.
Part III. The influence of external factors on
electrochemically active mixed culture biofilms
CHAPTER 7
The investigation of environmental parameters that affect the formation and performance of
electroactive biofilms stimulated this study since the majority of studies are restricted to a
neutral pH. Specifically how the pH value influences the biofilm growth (lag-time), steady state
anodic bioelectrocatalytic activity and microbial composition. I was involved in the replication
of fed-batch experiments at pH 6, 7 and 9 and also in the operation of continuous flow
experiments for pH-regime and buffer capacity studies.
CHAPTER 8
Most of the experiments designed to study electroactive biofilms in BESs are generally carried
out with one substrate or one microbial inoculum varying different operational parameters.
Therefore in order to exclude the influence of operational variables and to investigate only the
effect of an individual microbial inoculum source and an individual substrate, the experiments
here presented were conducted with half-cell set-ups under potentiostatic control with multiple
inocula and substrates. I was deeply involved in the recollection of inocula samples, preparation
of materials needed for half-cell experiments, later in the growth and maintenance of
electroactive biofilms and as well as in data collection, analysis and interpretation.
-26-
1.9 Comprehensive summary
Shewanella is frequently used as a model microorganism for microbial bioelectrochemical
systems (BESs) such as microbial fuel cells (MFCs) or microbial electrolysis cells (MECs). In
chapter 2, we used cyclic voltammetry (CV) to investigate extracellular electron transfer
mechanisms from Shewanella oneidensis MR-1 (WT) and five deletion mutants: membrane
bound cytochrome (ΔmtrC/ΔomcA), transmembrane pili (ΔpilM-Q, ΔmshH-Q, and ΔpilM-
Q/ΔmshH-Q) and flagella (Δflg). We demonstrate that the formal potentials of mediated and
direct electron transfer sites of the derived biofilms can be gained from CVs of the respective
biofilms recorded at bioelectrocatlytic (i.e. turnover) and lactate depleted (i.e. nonturnover)
conditions. As the biofilms possess only a limited bioelectrocatalytic activity, an advanced data
processing procedure, using the open-source software SOAS, was applied. The obtained results
indicate that S. oneidensis mutants used in this study are able to bypass hindered direct electron
transfer by alternative redox proteins as well as self-mediated pathways.
Figure: How does Shewanella transfer its electrons to solid acceptors? Using cyclic
voltammetry direct and mediated electron transfer of S. oneidensis MR-1 and related mutants
were investigated. The subsequent analysis, based on an elaborate open source software data -
processing, indicates a correlation of the maximum current density (x-axes of the graph) of the
respective mutant and its mediated electron-transfer ability (respective CV- peak height on the
y-axes).
-27-
It has been shown for anodic biofilms in MFCs that the microorganisms therein can be
influenced by the applied electrode potential. In chapter 3, we studied the influence of the
applied electrode potential on the anodic current production of Shewanella putrefaciens NCTC
10695. Furthermore, we used cyclic voltammetry (CV) and confocal laser scanning microscopy
(CLSM) to investigate the microbial electron transfer and biofilm formation. It is shown that the
chronoamperometric current density is increasing with increasing electrode potential from 3 µA
cm-2
at -0.1 V up to ~12 µA cm-2
at +0.4 V (vs. Ag/ AgCl), which is accompanied by an
increasing amount of biomass deposited on the electrode. By means of cyclic voltammetry we
demonstrate that direct electron transfer (DET) is dominating and the planktonic cells play only
a minor role.
Figure: Is the current generation, jmax, a function of the applied electrode potential?
Representative chronoamperometric fed-batch cycles of S. putrefaciens at graphite electrodes;
applied potentials: -0.1, 0, +0.1, +0.2, +0.3 and +0.4 V vs. Ag/AgCl; CV measurements during
turn-over (A) and non turn-over (B) conditions respectively.
0 1 2 3 4 5 6 7 8
-2
0
2
4
6
8
10
12
14
16
+0.4
+0.3
+0.2
+0.1
0.0
-0.1
AAA
j ma
x/
A c
m-2
time/ days
BBB
-28-
Crucial for the functioning of bioelectrochemical systems (such as MFCs and MECs) is the
complex protein architecture responsible for the electron transfer (ET) across the
bacteria/electrode interface. The ET pathway involves several multiheme redox proteins denoted
as outer membrane cytochromes (OMCs). In chapter 4 these OMCs were studied by a
combination of surface-enhanced resonance Raman scattering (SERRS) spectroscopy and
electrochemistry. The experiments presented in chapter 4 were performed on microbial biofilms
of S. putrefaciens. These have shown that OMCs do not contribute significantly to the
heterogeneous ET across bacteria/electrode interface. These studies have been performed on
biofilms grown on nanostructured Ag electrodes at the poised potential of +50 mV (vs.
Ag/AgCl). Although these conditions allow the formation of a biofilm on the Ag electrode, they
may have a negative impact on the amount of OMCs expressed by the bacteria (see chapter 3).
In fact, optimal biofilm growth requires pore positive potentials. However, these conditions
cannot be met by the Ag substrate, which undergoes oxidation at potentials higher than +150
mV (vs. Ag/AgCl).
Figure: Electrochemical measurements (A) performed in combination with SERRS (B)
allowed to control and monitor the activity of the microbial biofilm. This chapter aims at
providing the first spectroelectrochemical characterization of microbial biofilms of a strain of
the Shewanellaceae family by probing (i) structural information about the OMCs, (ii) the
participation of the OMCs to the ET, and (iii) the influence of soluble redox mediators
competing with OMCs. The experiments presented here contributed to elucidate the
function/structure relationship of OMCs in living cells, providing unique insight into the ET
across the bacteria/electrode interface. The development of novel analytical strategies to
overcome this limitation is presently under evaluation in our groups.
-29-
In chapter 5 we exploited electroactive bacteria in bioelectrochemical systems like MFCs that
promise a great potential in the context of sustainable energy supply and handling. A major
challenge in this context is to increase the performance of such systems, a necessity for a future
success of this new technology. During the past decade the average current densities of biofilm
anodes have already increased significantly from milliampere per square metre level to between
7 and 10 A m-2
. In this study it is demonstrated that by using three-dimensional carbon fiber
electrodes prepared by electrospinning and solution blowing the bioelectrocatalytic anode
current density reaches values of up to 30 A m-2
, which represents the so far the highest reported
values for electroactive microbial biofilms.
Figure: How do electroactive bacteria benefit from 3D materials? A) Biocatalytic current
generation at a 3D porous carbon fiber produced by gas-assisted electrospinning (GES-CFM)
modified carbon electrode in a model semi-batch experiment. The GES-CFM electrode was
modified by a wastewater-derived secondary biofilm grown in a half-cell experiment under
potentiostatic control. The electrode potential was 0.2 V vs. Ag/ AgCl. B) Scanning electron
microscopic image of an electroactive biofilm grown at a high porosity three-dimensional
structure carbon felt GES-CFM. The excellent bioelectrocatalytic performance of this material is
attributed to a structure that provides a habitat for the growth of electroactive bacteria up to a
maximum density supplemented by efficient substrate supply through the open pore structure.
The interconnections between the individual fibers of the nonwoven allow the formation of
cross-linked three-dimensional biofilms that benefit from an optimum electron transfer and
conduction.
-30-
In chapter 6 layered carbon fiber mats have been prepared by layer-by-layer (LBL)
electrospinning of polyacrylonitrile onto thin natural cellulose paper and subsequent
carbonization. The layered carbon fiber mat (CFM) has been proven to be a promising BES
anode material for MFCs and MECs, allowing high density layered biofilm propagation and
thus high bioelectrocatalytic anodic current density. Thick and continuous layered biofilms were
grown on these layered-carbon nanofiber mats and generated high current densities from waste
water derived biofilms. This investigation also revealed that, if the gap between the layers
within the layered-carbon nanofiber mats can be further increased in order to allow ideal
nutrient availability, thick layered biofilms might grow in every layer of the entire layered-CFM
and much larger current densities would be obtained. In summary, the cellulose-based carbon
fiber mat provides a low cost and highly efficient material for bioelectrocatalytic anodes in
microbial fuel cells.
Figure: Layered architecture of anode materials promotes the growth of electroactive
biofilms. A) Chronoamperometric (0.2 V vs. Ag/ AgCl) Biocatalytic current generation curves
from waste water derived biofilms of carbon fiber mats in a half-cell experiment; and B) SEM
image of a thick and continuous layered biofilm grown in the layered-CFM. In summary, here it
has been showed that small fiber diameter and proper pore size combined with sufficient three -
dimensionality- are essential features for the growth of high performance thick and continuous
biofilms.
-31-
It has been assumed that the treatment of wastewater could be one of the most appealing
applications for some BESs such as MFCs. Therefore, in order to make MFCs a successful
technology in wastewater treatment, researchers have to pay special attention to the
environmental and external factors that influence the biofilm. In chapter 7 the pH-value played a
crucial role for the development and current production of anodic microbial electroactive
biofilms. It was demonstrated that only a narrow pH-window, ranging from pH 6 to pH 9, was
suitable for growth and operation of biofilms derived from pH-neutral wastewater. Any stronger
deviation from pH neutral conditions led to a substantial decrease in the biofilm performance.
Thus, average current densities of 151 µA cm-2
, 821 µA cm-2
and 730 µA cm-2
were measured
for anode biofilms grown and operated at pH 6, 7 and 9 respectively. The microbial diversity of
the anode chamber community during the biofilm selection process was studied using the low
cost method flow-cytometry. Thereby, it was demonstrated that the pH value as well as the
microbial inocula had an impact on the resulting anode community structure. As shown by
cyclic voltammetry the electron transfer thermodynamics of the biofilms was strongly
depending on the solution’s pH-value.
Figure: Effect of pH in the performance of potentiostatic fed-batch electro-active biofilms
derived from primary wastewater. The more the pH-value during biofilm formation and
operation deviates from the pH of the bacterial source (pH neutral wastewater), the lower and
less efficient its bioelectrocatalytic activity becomes.
-32-
In chapter 8 the investigation of environmental parameters that affect the formation and
performance of anodic electroactive biofilms in MFCs were studied. In order to exclude the
influence of operational variables and to investigate only the effect of an individual microbial
inoculum source and an individual substrate, the experiments were conducted with half-cell set-
ups under potentiostatic control. A significant difference in current generation was observed for
all bioelectrochemical set-ups. Acetate-fed-reactor with primary wastewater inoculum showed
the highest current density (558 ± 27 μA cm-2
), followed by lactate-fed-reactor with primary
waste water inoculum (460 ± 54 μA cm-2
). The high performance with primary wastewater for
the formation of bioelectroactive biofilms demonstrated its ability as efficient microbial
inoculum source. Cyclic voltammograms (CVs) of all biofilms indicated the different electro-
chemical behaviour with both substrates. Maturity of biofilms was confirmed from a constant
maximum of current density production and a non-changing CV shape after several semi-batch
cycles (only for biofilms enriched from primary wastewater). For turnover CVs of biofilms
enriched from primary wastewater and both substrates the formal potential of the active site was
about -260 mV vs. Ag/ AgCl (see Figure). This clearly indicates that the used inocula
considerably influenced the enrichment of electrochemically active bacteria. For non-turnover
CVs, the electrochemical characterization of the biofilms reveals a strong similarity to
Geobacter sulfurreducens biofilms, which may indicate a dominating role of this bacterium in
the biofilms enriched from primary wastewater as source of inoculum.
Figure: Influence of the inoculum and substrate on the formation of electro-active
biofilms. Here exemplary CVs are shown for biofilms derived from primary wastewater set-ups,
the only experimental set-ups with a constant CV shape after the third semi-batch cycle.
-33-
CHAPTER II
2 Cyclic voltammetric analysis of the electron transfer of
Shewanella oneidensis MR-1 and nanofilament and
cytochrome knock-out mutants
2.1 Introduction
Electrochemically active bacteria (EAB) can transfer electrons to solid terminal electron
acceptors such as Fe(III), Mn(III), Cr(VI), and even to carbon electrodes in microbial fuel cells
(MFCs) (Chang, et al., 2006). These bacteria not only play a key role in nature's oxidation–
reduction cycles (Nielsen, et al., 2010) but also are the key component of microbial
bioelectrochemical systems (BES) (Rabaey, et al., 2010) (for an example on a BES see Fig. S9-1
in Supplementary information for Chapter II). Thus, the elucidation of the different microbial
electron transfer pathways is of fundamental interest as well as technological relevance.
Up to now, several classes of extracellular electron transfer mechanisms have been elucidated
for a wide range of microorganisms (Logan, 2009, Schröder, 2007). In principle, direct electron
transfer (DET) and mediated electron transfer (MET) can be distinguished. Whereas DET relies
on the physical contact of the redox active bacterial moiety, including redox proteins like
cytochromes (Busalmen, et al., 2008, Wigginton, et al., 2007) or bacterial nanowires (Gorby, et
al., 2006, Reguera, et al., 2005), with the solid terminal electron acceptor (e.g. iron (III) mineral
or anode of a BES), no such direct contact is necessary for MET. With MET, a dissolved
chemical compound that can serve as electron shuttle, i.e. mediator, facilitates the electron
transfer (Marsili, et al., 2008a, Rabaey, et al., 2005).
A wealth of different microorganisms, including single strain cultures as well as mixed
consortia, have been shown to be electrochemically active (Logan, 2009, Rabaey, et al., 2007)
and thus the analysis of the electron transfer pathways is actively being investigated. The
predominant model organisms studied are from the families Geobacteraceae (Lovley, 2008b)
and Shewanellaceae (Nealson and Scott, 2006). Whereas the electrochemical analysis of the
DET for Geobacter is well established (Fricke, et al., 2008, Marsili, et al., 2010, Srikanth, et al.,
2008), the latter microbe has been shown to possess more complex bioelectrochemical behavior
and the ability to generate sustained power in MFCs, even under constant exposure to dissolved
-34-
oxygen. In a recent study by Baron et al., cyclic voltammetry (CV) was used to demonstrate that
there are two redox active centres in adsorbed cells of Shewanella oneidensis; one responsible
for DET and one responsible for MET (Baron, et al., 2009). Furthermore, the authors showed
that the addition of exogenous flavins enhances mediated electron transfer. Recently, S.
oneidensis wild type and several electron transfer relevant mutants have been shown to differ
significantly concerning their bioelectrochemical activity (Bouhenni, et al., 2010).
Therefore, the aim of this study was to investigate the electron transfer properties of the
respective mutants within biofilms which were grown on an active electrode (i.e. in situ) using
cyclic voltammetry. A summary on electron transfer pathways for S. oneidensis and the
respective mutants used within this study are illustrated in the following section.
2.1.1 Extracellular electron transfer mechanisms of S. oneidensis MR-1 wild type and
mutants
The literature describing electron transfer mechanismsof S. oneidensis provides a complex
picture (see e.g. (Beliaev, et al., 2005, Bouhenni, et al., 2010, Gorby, et al., 2006, Hartshorne, et
al., 2007)) and is summarized in Fig. 2-1.
2.1.1.1 Direct electron transfer (DET)
For the facultative anaerobe S. oneidensis MR-1, there are two decahemes c-type cytochromes
(MtrC and OmcA) which are exposed on the outer cell surface and are assumed to be key
proteins for DET (Eggleston, et al., 2008, Firer-Sherwood, et al., 2008b, Fredrickson, et al.,
2008, Meitl, et al., 2009, Xiong, et al., 2006), yet are also reported to possess only limited rate
constants for electron transfer compared to mediator molecules (Baron, et al., 2009, Peng, et al.,
2010b).
Furthermore, as depicted in Fig. 2-1A these proteins can play a role in the mediated electron
transfer (MET) as the mediator reduction may take place on the outer cell surface. According to
the present models, this cytochrome facilitated DET and MET can be performed by the wild
type (WT) and all mutants used within this study, except ΔmtrC/ΔomcA due to a lack of outer
surface cytochromes. In addition to the cytochrome based electron transfer, pili outer surface
structures may play a role in extracellular electron transport. As shown in Fig. 2-1B and C, these
pili are believed to contribute to the microbial DET and MET.
-35-
Figure 2-1 Direct (DET) and mediated (MET) electron transfer pathways utilized by S.
oneidensis wild type and mutants. In every scheme it is indicated which strains can perform the
respective electron transfer mechanisms (Chang, et al., 2006, Nielsen, et al., 2010, Rabaey, et
al., 2010). A) Electron transfer via the cytochrome pool. Transmembrane pilus electron transfer
via B) pil-type pilus and via C) msh-type pilus, and D) biofilm formation behaviour. OM: Outer
membrane and IM: Inner membrane.
-36-
Furthermore, these pili are proposed to play a key role in the microbial cell attachment and thus
biofilm formation. Previous studies find that the three pili biogenesis knock-out mutants used
within this study (ΔpilM-Q, ΔmshH-Q and ΔpilM-Q/ΔmshH-Q) as well as the flagellin knock-
out mutant (Δflg) possess reduced biofilm forming ability (see Fig. 2-1D) (Bouhenni, et al.,
2010, Thormann, et al., 2004).
2.1.1.2 Mediated electron transfer (MET)
Several strains of the genus Shewanella can biosynthesize mediators to facilitate electron
transfer to the terminal electron acceptor. These mediators have been shown to be based on
flavins (e.g. (Biffinger, et al., 2010, Bouhenni, et al., 2010, Marsili, et al., 2008a, von Canstein,
et al., 2008)). For S. oneidensis, flavinmononucleotide (FMN) and riboflavin have been shown
to be redox shuttles in aerobic as well as anaerobic conditions (Velasquez-Orta, et al., 2010, von
Canstein, et al., 2008). In principle, a mediator molecule can be reduced in the inner cell or outer
cell membrane surface structures (Fig. 2-1A to C).
2.2 Materials and methods
2.2.1 General conditions
All chemicals were of analytical or biochemical grade and were purchased from Sigma-Aldrich
and Merck. If not stated otherwise, all potentials provided in this article refer to the Ag/AgCl
reference electrode (sat. KCl, 0.195 V vs. SHE). All microbial experiments were performed
under strictly sterile conditions.
2.2.2 Cell cultures and media
Sterilized LB Broth (Ringeisen, et al., 2006), and minimal media (Biffinger, et al., 2008) for
liquid cultures and LB/agar (Merck KGaA, Germany) plates were used for culture maintenance.
Single colonies of S. oneidensis MR-1 wild type and mutants (ΔmtrC/ΔomcA, ΔpilM-Q,
ΔmshH-Q, ΔpilM-Q/ΔmshH-Q and Δflg; obtained as reported elsewhere (Bouhenni, et al.,
2010)) were transferred to 15 mL of LB broth and incubated aerobically at room temperature
while shaking at 100 rpm (Universal shaker SM 30 A, Edmund Bühler GmbH, Germany) for 48
h. Afterwards, 15 mL of minimal media were added. Then 5 mL was used for subcultures in
growth medium (18 mmol/L Sodium lactate, PIPES Buffer 15.1 g/L; NaOH 3.0 g/L; NH4Cl 1.5
g/L; KCl 0.1 g/L; NaH2PO4∙H2O 0.6 g/L; NaCl 5.8 g/L; Mineral solution 10 mL/L (Atlas,
1993); Vitamin solution 10 g/L (Atlas, 1993); Amino acid solution 10 g/L (Bretschger, et al.,
2007)).
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2.2.3 Bioelectrochemical experiments
The bioelectrochemical experiments were carried out under potentiostatic control
(Potentiostat/Galvanostat VMP3, BioLogic Science Instruments, France) utilizing a three-
electrode arrangement, with a carbon rod working electrode (2.5 cm height×1.0 cm diameter,
CP-Graphite GmbH, Germany), a Ag/AgCl reference electrode (sat. KCl, Sensortechnik
Meinsberg, Germany) and a carbon rod (4.5 cm high×1.0 cm diameter) as counter electrode -
shielded by a 117 Nafion membrane. Sealed vessels (250 mL) served as the electrochemical cell
hosting the three-electrode arrangement (see Fig. S9-2 in Supplementary information). The
biofilm growth was performed in semi-batch chronoamperametric experiments at +0.2 V with
regular media replacement (addition of 200 mL fresh media solution to the 50 mL in the cell).
During these potentiostatic biofilm growth experiments aerobic conditions were assured
(Rosenbaum, et al., 2010a, von Canstein, et al., 2008) by pumping filtered air in the cells using
one fish pump (Elite air pump 799, Rolf C. Hagen Corp., Mansfield, MA. 02048, USA) per six
cells. Fresh electron donor and nutrients were supplied about every 24 h by removing 200 mL of
culture and replacing with 200 mL of fresh growth media.
Cyclic voltammetry (CV) was recorded during turnover conditions (TC), i.e. at the
bioelectrocatalytic substrate consumption, and during non-turnover (NTC), i.e. substrate
deprived, conditions at a scan rate of 1 mV s−1
. All CV experiments were performed under
anoxic conditions which were achieved before every experiment by bubbling nitrogen for 15
min in the solution. The headspace of the solution was also sparged with nitrogen during the
CV-measurement.
2.2.4 Data processing
Chronoamperometric maximum current densities (calculated per projected electrode surface
area) during at least 18 semi-batch cycles for established microbial biofilms for 3 independent
biofilm replicates were analyzed (see Figure S9-3 in Supplementary information). The standard
deviations are presented in Table 2-1. For in-depth data analyses of the cyclic voltammograms,
the open-source software SOAS (Fourmond, et al., 2009) was used for baseline (capacitive
current) correction for non-turnover conditions. Furthermore by using this software, the non-
turnover data was subtracted from the respective turnover data, and first derivatives were
calculated. Here all data are based on experiments during at least 18 semi-batch cycles of the 3
independent biofilm replicates, and the standard deviations are presented in Fig. 2-4.
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2.3 Results and discussion
2.3.1 Bioelectrochemical current production
Table 2-1 summarizes the maximum chronoamperometric current densities, derived from semi-
batch chronoamperometric experiments at 0.2 V vs. Ag/AgCl, for biofilms grown under aerobic
conditions of the S. oneidensis WT and the five mutant biofilms under investigation using
18mmoL/L lactate as the electron donor.
Table 2-1 Summary of the studied mutants and the achieved maximum current densities per
projected electrode surface area, the literature data are the reported maximum current densities
in MFC experiments at constant resistances.
Strain Mutant description
jmax / µA cm-2
This
worka
Ref.
Ab
Ref.
Bc
Ref.
Cd
Ref.
De
∆flg Flagella deletion mutant 9.5 ± 2.0 8 - - -
Wild Type S. oneidensis MR-1 wild-type,
ATCC 700550 7.9 ± 1.5 5 13 3.4 6
ΔpilM-Q Type IV pilus deletion mutant 7.7 ± 3.7 7.6 - - -
∆mtrc/
∆omcA
Outer membrane decaheme
c-type cytochromes MtrC and
OmcA deletion mutant
4.3 ± 0.8 0.6 2 0.7 <1
ΔmshH-Q Mannose-sensitive hemagglutinin
pilus deletion mutant 3.6 ± 1.9 3.2 - - -
ΔpilM-Q/
ΔmshH-Q
Type IV pilus and mannose-sensitive
hemagglutinin pilus deletion mutant 1.4 ± 1.6 2.2 - - -
aAverage data from chronoamperometric experiments at 0.2 V vs. Ag/AgCl calculated as
described in Section 2.4 and its respective standard deviation.bMFC experiments at an external
resistance of 100 kΩ after 200 h. c
MFC experiments at an external resistance of 10 Ω. d
MFC
experiments at an external resistance of 10 Ω. eChronoamperometric experiments at 0.043 V vs.
Ag/AgCl. Ref. A: (Bouhenni, et al., 2010); Ref. B: (Bretschger, et al., 2007); Ref. C: (Gorby, et
al., 2006); Ref. D: (Coursolle, et al., 2010).
All chronoamperometric biofilm growth experiments were made under oxygen exposure as
previous reports have shown that S. oneidensis WT preferentially forms thicker biofilms under
air-exposed conditions (Biffinger, et al., 2009). The growth pattern suits well with previous
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reports at other electrode potentials (e.g., 0.0 V vs. SCE (Peng, et al., 2010a) or 0.0 to 0.5 V vs.
Ag/AgCl (Peng, et al., 2010a)) and microbial activity was maintained for more than 18 days. For
comparison, Table 2-1 shows the current densities of the previous study on the respective
mutants under non-agitated air-exposed conditions in a microbial fuel cell study (Bouhenni, et
al., 2010, Bretschger, et al., 2007, Coursolle, et al., 2010, Gorby, et al., 2006). When further
comparing the WT current densities, the achieved maximum current density is well within the
expected range (Rosenbaum, et al., 2010a) - see Table S9-1 in Supplementary Information. The
focus of our analysis was the respective electron transfer mechanisms using cyclic voltammetry
and not maximizing current output as would be typical for MFC research.
2.3.2 Cyclic voltammetric analysis and data processing
In order to analyze the extracellular electron transfer mechanisms, cyclic voltammetry was
performed for all microbial biofilms/ suspensions during the growth cycles at maximum current
density (i.e. turnover) and at substrate depletion (i.e. non-turnover). Representative CVs for both
conditions are shown in Fig. 2-2A and B (nonturnover) and Fig. 2-3A and B (turnover).
Figure 2-2 A) and B) CVs for non-turnover conditions for S. oneidensis WT and mutants using
a scan rate of 1 mV s−1
; C and D) provide the respective baseline corrected curves.
-40-
Noteworthy, to avoid the oxygen disturbance on the CV measurements all experiments were
performed under strictly anoxic conditions. Furthermore, it has to be mentioned that initial
experiments where all bacteria were grown under anaerobic conditions did not yield sustainable
chronoamperometric and CV measurements.
Figure 2-3 A) and B) CVs for turnover conditions for S. oneidensis WT and mutants using a
scan rate of 1 mV s−1
.
As the CVs for both conditions do not show easily interpretable voltammetry (as it is the case
for e.g. Geobacter), the analysis of potential and actual electron transfer sites is not
straightforward (Fricke, et al., 2008). This behaviour can be attributed to the fact that the CV
current densities for turnover and non-turnover conditions vary less than one order of
magnitude, which indicates a rather small catalytic effect and hence low bioelectrocatalytic
activity of S. oneidensis. Noteworthy, the comparably high background current (that is due to
non-catalytically active, yet redox active microbial moieties and the exo-polysaccharide matrix)
in relation to the bioelectrocatalytic current densities might point towards i) a lower abundance
of redox centres per biomass and/ or ii) a lower turnover of the individual redox centre.
However, this question needs further investigation.
Thus, alternative, elaborate procedures for the CV analysis had to be followed. By using SOAS
(Fourmond, et al., 2009) baseline, i.e. capacitive current, corrected CVs for non-turnover
conditions could be generated - as shown in Fig. 2-2C and D. From these plots it can be
concluded that all studied bacterial strains use two different electron transfer pathways,
possessing formal potentials, Ef, of about -330 ± 45 mV (redox system I) and −70 ± 17 mV
(redox-system II), respectively.
-41-
Redox system I can be attributed to mediated electron transfer as its potential is in the range for
that found for soluble electron shuttles of Shewanellae in other studies (Biffinger, et al., 2008,
Marsili, et al., 2008a, Peng, et al., 2010b). However, the differing formal potential may be
attributed to the experimental conditions, e.g. free vs. biofilm-bound mediator and electrode
material, and at the electrode surface. Here a pH-decrease of the anodic reaction medium as well
as a pH-gradient near to the electrode surface (Torres, et al., 2008) will lead to a positive shift of
the formal potentials. In our potentiostatic set-ups, using shielded counter electrodes, we found
pH-values as low as pH 6 at the end of the growth cycle in the anodic compartment. This
influence of pH-microenvironment accounts as well for the membrane bound electron transfer
proteins, for which the redox-system II may be ascribed to a DET mechanism (Kim, et al.,
1999b).
The formal potentials of the surface proteins OmcA and MtrC varies within the literature. For
instance for a purified MtrC-protein, Ef-MtrC was found to be −270 mV vs. Ag/AgCl in pH 7
buffer solutions using basal plane graphite electrodes (Hartshorne, et al., 2007). Using Fe2O3
electrodes, Ef-OmcA was measured to be −210 mV when suspended in solution or −160 to −130
mV when adsorbed on the electrode surface (Eggleston, et al., 2008).
When studying whole cell suspensions of respective single knock-out mutants on haematite
electrodes, Ef-OmcA = −159 mV was identified (Meitl, et al., 2009). By measuring attached cells
on graphitic carbon electrodes, both formal potentials were estimated to be about −200 mV
(Baron, et al., 2009). Hence, these cytochromes possess a broad potential window of about 300
mV (Firer-Sherwood, et al., 2008b) and their actual formal potential seems strongly dependent
upon their microenvironment, and also here the pH-value direct at the electrode surface plays a
decisive role.
Furthermore, the potentially pilin-facilitated electron transfer mechanism was analyzed using the
Δflg, ΔpilM-Q, ΔmshH-Q, and ΔpilM-Q/ΔmsH-Q knock-out mutants. No discrimination from
the cytochrome-related signals was possible and thus no individual formal potentials of the pili-
related electron transfer could be identified, especially as no studies on the extracted
transmembrane moieties (ΔpilM-Q and ΔmshH-Q) are available. However, our data suggest that
the respective pili-related formal potentials are close to that of OmcA and MtrC, as all these
DET mechanisms are ultimately dependent on the same intracellular redox chains. This
assumption might furthermore explain why a DET signal was detected for ΔmtrC/ΔomcA that
may then be caused by the pili-related DET).
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In order to elucidate the reasons for the deviating bioelectrocatalytic activities of the S.
oneidensis WT and mutants - that could not be explained on differing numbers of active cells, an
elaborate analysis of the CV data for turnover (presented below) and non-turnover conditions
was performed. For non-turnover conditions, we analyzed the dependence of the maximum
bioelectrocatalytic activity on the peak properties of the baseline corrected CVs (Fig. 2-2C and
D). Here it was found that the peak height of the oxidation peak of the redoxsystem I, jpeak, was
a function of the chronoamperometric maximum current density, jmax (Table 2-1). This
dependency (Fig. 2-4) indicates that the concentration of the mediator molecule, which can be
assumed to correlate linearly with the respective height of the oxidation peak of redox-system I,
is required for the maximum current generation.
Figure 2-4 Plot of the base line corrected height of the oxidation peak of redox-system I (Δi−0.2)
as function of the maximum chronoamperometric current density of the respective microbial
culture.
Interestingly, this finding points towards an only inferior role of the DET mechanism for WT
and all mutants, which may sound logical when the low electron transfer rate constants of the
outer surface proteins mtrC and omcA are considered (Peng, et al., 2010b). Interestingly, for all
other peak areas and heights, such dependencies could not be identified (data not shown; finding
no correlation of jmax to the respective reduction peak of redox-system I has to be attributed the
-43-
limited CV-range, i.e. close vicinity of the signal to the vertex potential of the CV.). This is
contrary to our initial expectations that DET-related CV signal may be reduced for pilin-deletion
mutants. Yet, this finding might be explained as discussed below.
The analysis of the CVs for turnover conditions was not straightforward, as the calculation of
the derivatives did not yield a clear picture on the electron transfer sites. Therefore, we
performed the following analysis: The CVs for non-turnover conditions were subtracted from
the CVs for turnover conditions (Fourmond, et al., 2009), resulting in curves depicting the net-
catalytic current (Bard, et al., 2008). As Fig. 2-5 shows for the WT of S. oneidensis, two
catalytic waves could be identified. As the first half wave potential (approx. −300 mV) is very
similar to the formal potential of redox-system I - and thus found for flavin based mediators in
literature (Marsili, et al., 2008a) - it can clearly be attributed to MET. The ascription of the
second catalytic wave is more complex, as even its onset potential is more negative than the
formal potential of redox system II that was identified from non-turnover CVs (see above).
Figure 2-5 Plot of the corrected turnover CV signal and the performed analysis on the example
of S. oneidensis MR-1. (Similar plots of all strains can be found in Fig. S9-8 and Fig. S9-9 in the
Supplementary Information for Chapter 2).
Additionally, as can be clearly seen from Fig. 4-5, a half wave potential cannot be identified.
The latter finding may be attributed to several phenomena: First, the evolving proton gradient
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within the biofilm (Torres, et al., 2008) may lead to a (continuous) shift of the potential of the
electron transfer site. This finding is supported by recent studies investigating the pH
dependence of mixed culture biofilms (Patil, et al., 2011). Second, this result could be due to
phenomena that are well known from enzyme electrochemistry. Here analogue shaped catalytic
curves are ascribed to i) an inhomogenous coupling of the redox molecules to the electrode
surface and/or ii) slow electron transfer kinetics in relation to the substrate conversion kinetics
(Vincent, et al., 2007). Both factors are likely to play a role for whole cell biofilms. Noteworthy,
after smoothing and interpolation of the (non-turnover corrected) turnover CV curves, suitable
derivatives can be observed. For our measurements (Fig. S9-4 – Fig. S9-7 in the Supplementary
Information for Chapter 2) the maxima of these derivatives corresponds very well with the
formal potentials identified above.
Both catalytic waves can be found in all mutants, which indicate that in none of the microbial
cells was the DET completely inhibited, which is in line with our findings for non-turnover
conditions (see above). This result redox-signal when attached biofilms (Baron, et al., 2009) or
cell suspensions (Meitl, et al., 2009) were studied. However, when taking into account that S.
oneidensis possesses more than 42 (Meyer, et al., 2004) different cytochromes, of which 80%
are localized to the outer membrane (Heidelberg, et al., 2002), this finding might further support
the hypothesis that when one redox-protein is knocked out, other “bypass” molecules (including
other OMCs and/or pili) are used. In this context, two additional molecular biological studies
should be mentioned that illustrate the high versatility and complexity of S. oneidensis energy
metabolism. Kolker and co-workers revealed by a transcriptomics and proteomics based
analyses of 538 hypothetical genes, representing one third of the predicted number of proteins of
S. oneidensis MR-1 (Kolker, et al., 2005), that the respiratory versatility of this microorganism
may be explained not only by the high number of c-type cytochromes but also the existence of
numerous further, specialized genes that are directly related to the microbial energy conversion.
Beliaev et al. (Beliaev, et al., 2005), examined the globalmRNApatterns of S. oneidensis by
exposing the bacteria to different metal and non-metal electron acceptors. They have found that
only one of the 42 predicted c-type cytochromes (SO3300) displayed significantly elevated
transcript levels across all metal-reducing conditions. It has therefore been assumed that this
flavocytochrome c possessing a subunit with 4 hemes, participates significantly in the energy
metabolism and specifically possesses an outer surface electron transport role. Other mRNA-
levels, most prominently of MtrC and MtrA, were also slightly decreased in the presence of
metals.
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As an opportunity to (roughly) estimate the contribution of both electron transfer pathways (i.e.
DET and MET) to the bioelectrocatalytic activity of a respective microorganism, we propose the
following procedure (Fig. 2-5). The signal height of each catalytic wave is estimated at suitable
fixed potentials, where an almost full catalytic activity can be expected (here −0.2 V vs.
Ag/AgCl and 0.1 V vs. Ag/AgCl) and thus the share of each catalytic centre to the overall
maximum current can be estimated. The results of this analysis are summarized in Table 2-2.
The relative current share of redox-system I (Δi−0.2) is increased and consequently the share of
redox-system II (Δi+0.1) is decreased for all mutants (except ΔpilM-Q) in comparison to the wild
type. This finding indicates an increased contribution of mediated electron transfer when pili,
outer surface cytochromes or flagella are not available as DET pathways. However, the
differences, especially taking into account the standard deviations, are only minor for most of
the mutants — most pronounced with a MET share of 70% for motility inhibited mutant Δflg.
This finding corresponds well with the data presented in Fig. 2-4 and is an additional piece of
the complex extracellular electron transfer puzzle for Shewanella.
Table 2-2 Result of the CV subtraction analysis (details in Fig. 5 and the text).
Strain Relative share of the current/%
Redox-system I Redox-system II
ΔpilM-Q/ΔmshH-Q 57 ± 19 42 ± 16
ΔpilM-Q 48 ± 24 51 ± 43
Wild-type 52 ± 27 47 ± 16
ΔmshH-Q 61 ± 26 38 ± 14
Δflg 71 ± 19 28 ± 14
ΔmtrC/ΔomcA 62 ± 44 37 ± 17
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2.4 Conclusions
In this study we exploited cyclic voltammetry as a tool for the in situ study of S. oneidensis wild
type and mutant biofilms grown at electrode surfaces. Since the catalytic activity of these
microorganisms was found to be limited (as reflected by only low current densities) the analysis
of the electron pathways is not as straightforward as for bacteria that do not possess the ability to
self-mediate extracellular electron transfer (Fricke, et al., 2008). In order to analyze the CV data,
an elaborated data processing was performed. By this data analysis, the formal potentials of the
direct and mediated electron transfer were identified and the share of each electron transfer
pathway to the overall current production could be estimated. However, it was not possible to
elucidate thermodynamic/ mechanistic differences in the direct electron transfer for respective
knock-out mutants. Furthermore our results indicate that mutants possessing knock-outs for
potential DET-related proteins bypass this deficiency by alternative DET redox-carriers and
self-mediated pathways. Here, more sensitive electrochemical (e.g. square wave voltammetry),
spectroscopic and related techniques in combination with molecular biological approaches (e.g.
transcriptomics) need to be exploited in future research.
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CHAPTER III CHAPTER III
3 Study of Shewanella putrefaciens biofilms grown at
different applied potentials using cyclic voltammetry and
confocal laser scanning microscopy
3.1 Introduction
Although, the ability of microorganisms to create an electrochemical potential was discovered
more than 100 years ago (Potter, 1911), it was mainly during the last decade that this field of
research was developing most dramatically (Schröder, 2011), as it was found that the ability to
transfer electrons to extracellular electron acceptors is naturally occurring in several microbial
species. This interest thereby is triggered by fundamental inquisitiveness (Hernandez and
Newman, 2001, Schröder, 2007, Watanabe, et al., 2009) as well as by the development of
seminal sustainable technologies based on microbial extracellular electron transfer - also known
as microbial bioelectrochemical systems (BES) (Rabaey, 2010, Rabaey and Rozendal, 2010).
Up to now, several microorganisms have been studied in order to elucidate their specific
extracellular electron transfer mechanisms, including Lactococcus lactis (Masuda, et al., 2010),
Saccharomyces cerevisiae (Ducommun, et al., 2010), Pseudomonas sp. CMR12a (Pham, et al.,
2008), Hansenula anomala (Prasad, et al., 2007), Proteus vulgaris (Rawson, et al., 2011) and
Lyngbya sp. and Nostoc sp. (Pisciotta, et al., 2011). Here, the two families of Gram-negative
bacteria Shewanellaceae (Myers and Nealson, 1988, Nealson and Scott, 2006) and
Geobacteraceae (Caccavo, et al., 1994, Lovley, 2008b, Malvankar, et al., 2011, Richter, et al.,
2009) are the most widespread model organisms. Thereby, different electron transfer strategies
have been commonly described among the various species, which are summarized as follows: i)
direct electron transfer (DET) via membrane-bound redox-enzymes (Inoue, et al., 2011, Millo,
et al., 2011, Strycharz, et al., 2011) or via bacterial nanowires (El-Naggar, et al., 2010, Gorby, et
al., 2006, Malvankar, et al., 2011, Reguera, et al., 2005) and ii) mediated electron transfer
(MET) via redox shuttle substances, e.g. (Jiang, et al., 2010, Marsili, et al., 2008a), usually
secondary microbial metabolites.
-48-
For the family of Shewanellaceae – all being considered to be facultative anaerobes - several
members have been studied on their extra-cellular electron transfer behavior. Most prominently
S. oneidensis MR-1 (Baron, et al., 2009, Meitl, et al., 2009, Okamoto, et al., 2011, Sun, et al.,
2010), but also S. oneidensis MR-4 (Marsili, et al., 2008a), S. putrefaciens W3-18-1
(Bretschger, et al., 2010a), S. putrefaciens IR-1 (Kim, et al., 1999b, Kim, et al., 1999d, Kim, et
al., 2002), S. putrefaciens SR-21 (Kim, et al., 2002), S. loihica PV-4 (Bretschger, et al., 2010a,
Nakamura, et al., 2009a, Nakamura, et al., 2009b, Okamoto, et al., 2009, Zhao, et al., 2010b), S.
decolorationis NTOU1 (Li, et al., 2010, Li, et al., 2009a), S. japonica KMM 3299 (Biffinger, et
al., 2010), S. frigidimarina NCIMB400 (Pankhurst, et al., 2006, Turner, et al., 1999) and S.
marisflavi EP1 (Huang, et al., 2010). Commonly, it is assumed that the two decaheme c-type
cytochromes MtrC and OmcA, both facing the extracellular environment, play a key role in the
direct electron transfer mechanism (DET) (Eggleston, et al., 2008, Firer-Sherwood, et al.,
2008b, Fredrickson, et al., 2008, Hartshorne, et al., 2007, Meitl, et al., 2009, Xiong, et al.,
2006). Thereby, MtrC and OmcA, are part of a complex transmembrane cytochrome pool
involving more than 40 proteins (Beliaev, et al., 2005).
The MET of Shewanellaceae exploits redox-shuttles including humic substances (Hong, et al.,
2007), melanin (Turick, et al., 2009, Turick, et al., 2002), menaquinone (Lies, et al., 2005) as
well as riboflavin (von Canstein, et al., 2008), flavinmononucleotide (Biffinger, et al., 2010,
Velasquez-Orta, et al., 2010) and their derivatives. Further it is considered to depend on
intracellular electron transfer to the redox-shuttle as well as extracellular electron transfer by
MtrC or OmcA, e.g. (Biffinger, et al., 2010, Bouhenni, et al., 2010, Marsili, et al., 2008a).
Thereby it has been shown that some strains of the Shewanella family (e.g. S. oneidenis MR-1
(Jiao, et al., 2011, Marsili, et al., 2008a), S. loihica PV-4 (Newton, et al., 2009), S. baltica
Os155 and Os195 (von Canstein, et al., 2008), S. frigidimarina NCIMB400 (von Canstein, et al.,
2008) and S. decoloratioans NTOU1 (Li, et al., 2010)) can biosynthesize these mediators under
aerobic as well as anaerobic cultivation. However, the ability to synthesize suitable amounts of
these electron shuttles (especially for anaerobic conditions, where energy for biosynthesis is
limited) is not unequivocally established for all Shewanella species, e.g. (von Canstein, et al.,
2008). In this course it is of crucial importance to consider the exact conditions during biofilm
growth and development (Harnisch and Rabaey, 2012).
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3.1.1 Influence of the electrode potential on electroactive microbial biofilms
It has been shown for anodic mixed culture derived biofilms that the applied electrode potential
determines decisively the bacterial composition, e.g. (Torres, et al., 2009), of the gained biofilm.
However, also pure culture anodic biofilms and the microorganisms therein can be influenced
by the applied electrode potential. Cho and Ellington (Cho and Ellington, 2007) demonstrated
for S. oneidensis MR-1 that during chronoamperometric biofilm growth the lag-period
decreased from 90 days at 0 mV to 5 days at 500 mV (vs. Ag/AgCl), respectively with
increasing electrode potential; whereas the current density was almost constant. Furthermore,
Shewanella oneidensis MR-1 is believed to show some motility towards electrodes and thus the
availability of its cellular appendices (wires, pili and flagella) clearly determines its electron
transfer performance (Carmona-Martínez, et al., 2011). Recently Harris and colleagues (Harris,
et al., 2010) observed an increase in cell swimming speed when whole cells of Shewanella
species (S. oneidensis MR-1, S. amazonensis SB2B and S. putrefaciens CN32) were exposed to
varying electrode potentials. In addition, Liu and colleagues (Liu, et al., 2010a) investigated
microbial extracellular electron transfer activity of S. loihica PV-4 by applying different
electrode potentials (from -0.4 V to 0.2 V vs. Ag/ AgCl). Thereby, they demonstrated a clear
dependence of the activity on the applied electrode potential, with an activity increase for
potentials more positive than -220 mV. Furthermore, Liu et al. (Liu, et al., 2011) have recently
shown on the example of S. oneidenis MR-1 and S. loihica PV-4 that, depending on the
potential of bacteria cultivation, the electron transfer pathways can be switched from DET to
MET and the formal potential of the DET (Liu, et al., 2010a) is also triggered by the redox-
conditions during cultivation.
Within the context of correlating the applied electrode potential (i.e. potential of the microbial
terminal electron acceptor) and the bacterial activity, it was the aim of this study to elucidate the
electrochemical response of the S. putrefaciens NCTC 10695, a strain that is studied in BES for
the first time. For a comparison with other S. putrefaciens see table S10-2. Therefore, the
influence of the applied electrode potential on the electron transfer mechanisms was investigated
using cyclic voltammetry (CV) and biofilm morphology by means of confocal laser scanning
microscopy (CLSM). CLSM has recently been applied for the study of electroactive biofilms,
e.g. to monitor their pH-gradients (Babauta, et al., 2011), but has not yet been exploited for the
quantification of biofilm biomass for different electrode potentials.
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3.2 Materials and methods
3.2.1 General conditions
All chemicals were of analytical or biochemical grade and were purchased from Sigma-Aldrich
and Merck. If not stated otherwise, all potentials provided in this article refer to the Ag/AgCl
reference electrode (sat. KCl, 0.195 V vs. SHE). All microbial experiments were performed
under strictly sterile conditions.
3.2.2 Cell cultures and media
Sterilized LB broth (Ringeisen, et al., 2006), and minimal media (Bretschger, et al., 2007) (18
mmol/ L sodium lactate, PIPES buffer 15.1 g×L-1
; NaOH 3.0 g×L-1
; NH4Cl 1.5 g×L-1
; KCl 0.1
g×L-1
; NaH2PO4∙H2O 0.6 g×L-1
; NaCl 5.8 g×L-1
; Wolfe’s mineral solution 10 mL×L-1
(Atlas,
1993); Wolfe’s vitamin solution 10 mL×L-1
(Atlas, 1993); amino acid solution 10 mL×L-1
(Bretschger, et al., 2007)) for liquid cultures and LB/agar (Merck KGaA, Germany) plates were
used for culture maintenance.
Single colonies on LB/agar plates freshly streaked from a frozen glycerol stock culture
(Coursolle, et al., 2010) of S. putrefaciens wild-type NCTC 10695 (Nealson and Myers, 1992,
Pivnick, 1955, Venkateswaran, et al., 1999, Vogel, et al., 1997) (DSM No.: 1818, DSMZ -
German Collection of Microorganisms and Cell Cultures GmbH, Braunschweig, Germany);
were transferred to 15 mL of LB broth and incubated aerobically at room temperature while
shaking at 100 rpm (Universal shaker SM 30 A, Edmund Bühler GmbH, Germany) for 48 h
(Carmona-Martínez, et al., 2011). Afterwards 15 mL of culture were spun down at 3000 rpm
during 10 min (Heraeus Megafuge 1.0, Germany). The pellet was resuspended in 15 mL of
minimal media and transferred to a sterile Erlenmeyer flask with 185 mL of minimal media for
72 h of cultivation using the same conditions. Finally 200 mL of media were centrifugated at
3000 rpm during 10 min, the pellet was resuspended in 15 mL of minimal media and injected in
the electrochemical cell.
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3.2.3 Bioelectrochemical set-up and experiments
Polycrystalline carbon rod (CP Graphite GmbH, Germany) was used as working (2.0 cm height
x 1.0 cm diameter) and as counter (7.0 cm height x 1.0 cm diameter) electrodes for the growth
and investigation of the (anodic) electrocatalytic biofilms. Carbon electrodes were glued with
stainless steel wire (AISI 304, Fe/Cr18/Ni10, Goodfellow GmbH, Nauheim, Germany) using a
two-component resin (Epoxy resin HT 2 + Hardener HT 2, HY-POXY® Systems, SC, USA)
mixed with carbon black particles (Vulcan XC-72, Cabot Corporation GMBH, Frankfurt am
Main, Germany).
All bioelectrochemical experiments were conducted under strictly sterile conditions and under
potentiostatic control, using Ag/AgCl reference electrode (sat. KCl, 0.195 V vs. SHE,
Sensortechnik Meinsberg, Germany). S. putrefaciens biofilms were grown in potentiostatic half-
cell experiments at six different anode potentials (-0.1, 0, +0.1, +0.2, +0.3 and +0.4 V) at 30 °C
(Owen, et al., 1978) – see also Figure S10-1. To assure comparability and reproducibility up to
six electrodes were measured simultaneously in one electrochemical cell using an Autolab 30
potentiostat (Ecochemie, The Netherlands) equipped with six array channels. The biofilm
growth was performed in semi-batch chronoamperametric experiments at different applied
potentials with a regular Shewanella cells addition procedure (Baron, et al., 2009) and media
replacement as described below. For these potentiostatic biofilm growth experiments, initial
aerobic conditions were assured according to previous reports (Biffinger, et al., 2009).
Therefore, before adding the growth media into the electrochemical cell (Rosenbaum, et al.,
2010a, von Canstein, et al., 2008) filtered air was pumped into the fresh minimal media using a
fish pump (Elite air pump 799, Rolf C. Hagen Corp., MA, USA) per 400 mL for more than 1 h.
Fresh electron donor and nutrients were supplied about every 72 h by removing 320 mL of
culture (representing 80% of the cell volume) and replacing with fresh minimal media and fresh
Shewanella cells according to Section 3.2.2. Cyclic voltammograms were recorded for turnover
and non-turnover conditions according to (Carmona-Martínez, et al., 2011).
3.2.4 Electrochemical data processing
All data are based on experiments of 3 independent biofilm replicates and the standard
deviations are presented. The maximum current density, jmax, (calculated per projected electrode
surface area) of a batch-cycle was taken as a measure of activity and was analyzed during at
least 6 semi-batch cycles for established microbial biofilms. For in-depth data analyses of the
cyclic voltammograms the open–source software SOAS (Fourmond, et al., 2009) was used for
baseline (capacitive current) correction for non-turnover conditions.
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3.2.5 Confocal Laser Scanning Microscopy
Shewanella biofilms on polycrystalline graphite electrodes were examined by CLSM after
staining with nucleic acid-specific fluorochromes. For this purpose, whole cylindrical electrodes
(~2.0 cm in exposed length) were mounted on a plastic Petri dish with silicon glue, subsequently
stained with Syto9 (Molecular Probes, OR, USA) and incubated at room temperature for 5 min
(Neu, et al., 2010). The laser microscope (SP5X, Leica Germany) was equipped with a super
continuum light source and an upright microscope. The system was controlled by the LEICA
CONFOCAL software version 2.4.1 Build 1537 (Leica, Germany). For imaging, the stained
biofilms were covered with tap water and examined with a 63x 0.9 NA objective lens. Each
Shewanella biofilm attached to the electrode was scanned at ten different locations from top to
bottom. The settings for excitation and emission/detection were as follows: excitation with the
white laser at 483 nm, detection of reflection from 475 to 495 nm and of Syto9 from 500 to 570
nm. For recording the bacterial emission signal, the lookup table "glow-over-under" was used in
order to optimally adjust signal to noise ratio and taking advantage of the full dynamic range of
the photomultiplier. Quantification was done with an extended version of ImageJ
(rsbweb.nih.gov.ij) as described elsewhere (Staudt, et al., 2004). The 8 bit data sets were
thresholded at 60. Image data sets were printed from Photoshop (Adobe).
3.3 Results and discussion
3.3.1 Bioelectrochemical current production
Figure 3-1 shows for exemplary fed-batch cycles the chronoamperometric (CA) current
production of S. putrefaciens NCTC 10695 for different applied electrode potentials. As can be
seen the maximum current density, jmax, as well as the transferred charge of each electrode
differed significantly. Since all electrodes were hosted within one vessel in order to assure
similar biological conditions for all electrodes in each experiment (see material and methods),
no coulombic efficiencies can be calculated for the individual electrode potentials.
However, when taking jmax at a given electrode potential a clear trend can be observed. As
Figure 3-2 shows the current density increases from less than 3 µA cm-2
at -0.1V up to ~12 µA
cm-2
at +0.4 V.
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Figure 3-1 Representative chronoamperometric fed-batch cycles of S. putrefaciens at graphite
electrodes; applied potentials: -0.1, 0, +0.1, +0.2, +0.3 and +0.4 V vs. Ag/AgCl; CV
measurements during turn-over (A) and non turn-over (B) conditions respectively.
Figure 3-2 Chronoamperometric current density of S. putrefaciens as function of the applied
electrode potential.
0 1 2 3 4 5 6 7 8
-2
0
2
4
6
8
10
12
14
16
+0.4
+0.3
+0.2
+0.1
0.0
-0.1
AAA
j ma
x/
A c
m-2
time/ days
BBB
-0.1 0.0 0.1 0.2 0.3 0.40
3
6
9
12
15
j ma
x/
A c
m-2
applied potential/ V vs. Ag/AgCl
-54-
Thus, the measured current densities are well in line with previous studies on Shewanellaceae
using plain carbon electrodes (see Table S10-1 in SI for a comparative data compilation). No
biofilms could be seen by the naked eye on all electrodes, but the solution showed significant
turbidity and a reddish pellet of bacteria was formed when spinning it down (see Fig. S1-inset).
This finding is well in accordance with previous reports on Shewanaellaceae (Babauta, et al.,
2011, Coursolle, et al., 2010, Okamoto, et al., 2011, Yang, et al., 2011), yet in contrast to pure
culture biofilms of Geobacter species or mixed culture derived biofilms dominated by
Geobacteraceae. In order to elucidate reasons for the varying electrochemical performance at
the different electrode potentials, cyclic voltammetry (CV) and confocal laser scanning
microscopy (CLSM) were employed.
3.3.2 Cyclic voltammetric analysis
Figure 3-3 shows the CVs of the electrodes under turn-over conditions, i.e. in the presence of the
electron donor lactate. All CVs show an increasing bioelectrocatalytic activity for all potentials
more positive than 0.0V. All CVs possess a half wave potential of about ~10 mV (as can be
derived from the maxima of the respective first derivatives (Fricke, et al., 2008, Marsili, et al.,
2008b)). This potential can commonly be ascribed to DET-proteins (see e.g. (Hartshorne, et al.,
2007, Shi, et al., 2006)). In this course it has to be noticed that this is the first voltammetric
study of S. putrefaciens NCTC 10695 (for S. putrefaciens from other sources see Table S10-2).
Thus, the results for turn-over conditions point out that the MET via soluble mediators plays
only a minor role, as the formal potentials, Ef, for these electron shuttling compounds (and thus
the corresponding half wave potential of the turn-over CV) are more negative. Noteworthy, the
riboflavin concentration of our minimal media nutrient broth was only about 0.1 μM (see
materials and methods). When this concentration was increased to 1 μM, the turn-over CV-
shape changed significantly and clearly illustrated that the riboflavin based MET was dominant
in these conditions (see SI Figures S10-2 and S10-3). Thus, as no significant MET associated
bioelectrocatalysis was found after more than two weeks of continuous cultivation, our results
suggest that S. putrefaciens NCTC 10695 did not synthesize relevant amounts of electron shuttle
compounds. This finding is in contrast to other species of the Shewanella family like S.
oneidensis MR-1 (Jiao, et al., 2011, Marsili, et al., 2008a, von Canstein, et al., 2008). We
suggest to ascribe this finding to the particular growth conditions especially i) the almost anoxic
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behavior and ii) low mediator concentration in the medium and thus fast establishment of a
biofilm at the electrode.
Figure 3-3 A) Representative cyclic voltammograms of S. putrefaciens for turn-over conditions
and B) respective first derivatives of the voltammetric curves; scan rate: 1 mV s-1
.
In order to study the biofilm associated direct electron transfer processes in more detail, CVs for
non turn-over (lactate depleted) conditions were recorded (see Figure 3-4 A for the raw data and
Figure 3-4 B for the baseline corrected traces). It can be clearly seen that the non turn-over CVs
for the selected electrode potentials differed significantly. As all electrodes were hosted within
one vessel, these differences cannot be attributed to the suspended microbial cells, but to the
individual biofilms. Noteworthy, no CV signals were detected for electrodes poised at -0.1V and
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0.0V during biofilm growth, which can clearly be attributed to the absence of the interaction
with bacteria on the electrode surface (vide infra).
As Figure 3-4 shows, the CV analysis of all further electrodes revealed that the formal potential
of the redox-active species is similar for all growth potentials (about -60 mV, see also Table
S10-3). This value is close to the maximum in the first derivatives of the turn-over CVs (Figure
3-3) showing that the detected redox couple is responsible for the bioelectrocatalytic electron
transfer. As can be seen from Figure 3-4 the peak heights and areas of the oxidation and
reduction peaks are increasing with more positive electrode potential during biofilm growth.
Figure 3-4 A) Cyclic voltammograms for non turn-over conditions for S. putrefaciens using a
scan rate of 1 mV s−1
; B provides the respective baseline corrected curves.
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Deriving from the data exemplary shown in Figure 3-4, Figure 3-5 depicts the peak areas and
peak heights of the respective oxidation and reduction peak as function of the applied potential
during CA. As the peak area can be considered to be a measure of the amount of a redox species
on the electrode surface (Wang, et al., 2002a, Wang and Wang, 2004), this analysis shows that
with increasingly positive potential during bacterial growth more DET-protein is deposited on
the electrode surface. As the increasing amount of DET protein can be either attributed to an
increasing cell number or to an increase in the protein amount per individual cell confocal laser
scanning microscopy (CLSM) was exploited to study these biofilms at the morphological level.
Figure 3-5 Plot of the base line corrected height (○) and area (□) of the oxidation and reduction
peaks of redox-system shown in Fig. 3-4 as function of the applied potential. For visual
convenience, reduction peak areas are shown as negative values.
As already discussed above, the detected formal potential is close to but not identical to those of
MtrC and OmcA commonly reported in literature for different experimental conditions, e.g. on
isolated proteins (Hartshorne, et al., 2007). Here it should be noticed that additional cyclic
voltammetry experiments conducted on S. putrefaciens cells suspensions (data not shown) did
not provide redox peaks in any condition tested, i.e. 1) bacteria growing microaerophilically in
LB broth; 2) cells growing aerobically in minimal media (Bretschger, et al., 2007) and 3) cells
-0.1 0.0 0.1 0.2 0.3 0.4
-1
0
1
2
3
pea
k h
eig
ht/
A
cm
-2
applied potential / V vs. Ag/AgCl
-0.2
0.0
0.2
0.4
pea
k a
rea/
C
cm
-2
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growing anaerobically in (Miller and Wolin, 1974) minimal media (Bretschger, et al., 2007,
Marsili, et al., 2008a, Meitl, et al., 2009) using sodium fumarate as electron acceptor.
3.3.3 Biofilm imaging using confocal laser scanning microscopy (CLSM)
Figure 3-6 Maximum intensity projection of confocal laser scanning microscopy data sets
showing Shewanella putrefaciens biofilms grown on electrode surfaces at different applied
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potentials. A) -0.1 V, B) 0 V, C) +0.1 V, D) +0.2 V, E) +0.3 V and F) +0.4 V; (all vs. Ag/AgCl).
Colour allocation: reflection of electrode – grey, nucleic acid stained bacteria – green.
Figure 3-7 Biofilm quantification of Shewanella putrefaciens biofilms grown on electrode
surfaces at different applied potentials.
In order to study the biofilm formation at the electrode surface exemplary CLSM measurements
were performed. As Figure 3-6 shows the number of cells (Franks, et al., 2009) seems to
increase with more positive electrode potential. This is confirmed by a biomass quantification
based on the CLSM imaging and subsequent digitital image analysis (Figure 3-7). Please note
that the high numeric value in Figure 3-7 for -0.1V can only be attributed to an inhomogeneous
biofilm growth and biofilm sloughing. This effect needs further consideration in future studies
with focus on biofilm development over time. The data is showing a linear increase of biomass
with more positive electrode potential. Nevertheless the relatively high standard deviation is
obvious and well known from other studies, e.g. (Lewandowski, et al., 2004), indicating a non-
uniform (rough) biofilm coverage. Interestingly, as Figure 3-6 shows, no complete coverage of
the electrode surface and decent biofilm formation was detected for all applied electrode
potentials. This finding is well in line with previous studies on Shewanella (Babauta, et al.,
2011, Coursolle, et al., 2010, Okamoto, et al., 2011, Yang, et al., 2011), but in clear contrast to
Geobacter species (Franks, et al., 2009, Williams, et al., 2009).
-0.1 0.0 0.1 0.2 0.3 0.4
0
1000
2000
3000
4000
applied potential/ V vs. Ag/AgCl
bio
ma
ss/
m3
-60-
Therefore, and in combination with the dominance of DET and planktonic cell growth, one may
conclude that attachment of S. putrefaciens cells was not permanent, indicating an intermittent
cell-electrode contact for electron release (Harris, et al., 2010). Furthermore, this shows clearly
the attraction of the more positive potential for biofilm formation, which is in line with a report
on the bacterial movement to electrodes, termed “electrokinesis” (Harris, et al., 2010).
3.4 Conclusions
This study on the electroactive microorganism S. putrefaciens NCTC 10695 shows that:
The chronoamperometric current generation, jmax, is a function of the applied electrode
potential; it increases from 3 µA cm-2
at -0.1V up to ~12 µA cm-2
at +0.4 V.
The amount of biomass deposited on the electrode is a function of the electrode potential
and steadily increase by factor 5 from 0V to +0.4 V.
For the range of applied electrode potentials studied the direct electron transfer (DET) is
dominating.
The bioelectrocatalytic current generation, the CV signal of the DET protein and the
biomass coverage are clearly linked by the applied electrode potential during biofilm
growth.
Interestingly, for electrode potentials more positive than the formal potential of the active site (-
60mV) the current density and biofilm formation increase. The same phenomena is found for the
turn-over CVs not showing a plateau of current density, which is in contrast to species like
Geobacter (Fricke, et al., 2008).
Most of the results found for S. putrefaciens NCTC 10695 are well in line with literature,
however some vary. These variations might be attributed to i) the specific nature of the strain S.
putrefaciens NCTC 10695 or ii) the specific growth conditions and thus may change in other
environments, e.g. presence vs. absence of O2 in the medium or constant vs. varying electrode
potentials. Therefore, they are in a row of studies on Shewanella species, providing snapshots
but not allow drawing conclusive pictures. Thereby, this study highlights exemplary the need for
a unified framework of biofilm growth and operation of electroactive biofilms, like
Shewanellaceae (Harnisch and Rabaey, 2012) in order to extract more (universal) information
out of the gained data.
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CHAPTER IV CHAPTER IV
4 Spectroelectrochemical analysis of intact microbial
biofilms of Shewanella putrefaciens for sustainable energy
production
4.1 Introduction
The ability of some microbes to respire insoluble metal oxides and electrodes relays on the
peculiar extracellular electron transfer (ET) strategy they evolved. In fact this mechanism by
which microorganisms generate energy for cell growth and maintenance (Hernandez and
Newman, 2001), allows the bacteria to transfer electrons from their internal metabolism through
a chain of trans-membrane proteins to insoluble metal electron acceptors placed outside the
microbial cell (Fig. 4-1). In the early 1990s, environmental microbiologists came to realize the
importance of microbial ET of insoluble metal electron acceptors in several biogeochemical
cycles and progressively applied this extraordinary finding, e.g., on the bioremediation of
contaminated sites (Lovley, 1991, Nealson, et al., 1991). More recently this finding has been
used in an multidisciplinary way not only to study the fundamentals of microbial ET but also to
apply this concept in the so-called Bioelectrochemical systems (BESs) (Rabaey, 2010), e.g., for
the production of: i) electricity (Logan, et al., 2006), ii) hydrogen as a clean fuel (Logan, et al.,
2008) and iii) useful chemicals (Rabaey and Rozendal, 2010) such as hydrogen peroxide, among
other possible applications.
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Figure 4-1 Principle representation of a BES operating in the DET mode (see below). Electrons
derived from the oxidation of the organic substrate catalyzed by the bacterial cell are shuttled to
the electrode via OMCs.
The scientific information available on microbial ET has rapidly increased mainly due to the
discover of two model bacteria capable of reducing insoluble metal electron acceptors:
Shewanella oneidensis MR-1 (Myers and Nealson, 1988) and Geobacter metallireducens GS-15
(Lovley and Phillips, 1988). Whereas the electrochemical analysis of the ET mechanisms for
Geobacter is well established (Fricke, et al., 2008, Marsili, et al., 2010, Millo, et al., 2011,
Srikanth, et al., 2008), Shewanella has been shown to possess more complex bioelectrochemical
behavior.
Interestingly, Geobacter and Shewanella species rely upon different microbial ET mechanisms,
i.e. direct and a combination of direct-mediated ET, respectively (Bretschger, et al., 2010b,
Gralnick and Newman, 2007, Hernandez and Newman, 2001, Marsili and Zhang, 2010,
Schröder, 2007, Watanabe, et al., 2009). In direct electron transfer (DET), direct electrical
contact between the bacterial cell and the electrode is provided by outer membrane cytochromes
(OMCs). These multiheme redox proteins embedded within the outer membrane of the bacterial
cell shuttle electrons between the respiratory chain and the insoluble electron acceptor through a
densely packed chain of heme groups. Recently, the DET has been proposed to occur also
through cellular appendages facing the extracellular environment (i.e., microbial nanowires).
Although these appendages have been initially observed for Shewanellaceae (El-Naggar, et al.,
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2010, Gorby, et al., 2006), they have been recently found also for Geobacteraceae (Malvankar,
et al., 2011, Reguera, et al., 2005) families, where they have been proposed to sustain long-
range metallic-like ET (Malvankar et al., 2011). In mediated electron transfer (MET), the
bacteria release soluble redox mediators such as flavin or melanin, that go through a series of
reduction and oxidation processes taking place between the bacterial cell and an extracellular
insoluble compound (Turick, et al., 2002, von Canstein, et al., 2008).
It is worth noting that one mechanism does not exclude the other, in such a way that DET and
MET can function simultaneously within the same microbial community. This is the case of
Shewanella oneidensis MR-1. As recently reported by Shi and co-workers (Shi, et al., 2009) the
DET performed by S. oneidensis MR-1 depends on inner membrane cytochromes (IMCs) and
OMCs that are known to be directly involved in the reduction of insoluble metals that act as
extracellular electron acceptors (or in the case of BESs: electrode materials). These proteins
include the inner membrane tetrahaem c-Cyt CymA that is a homologue of NapC/NirT family of
quinol dehydrogenases, the periplasmic decahaem c-Cyt MtrA, the outer membrane protein
MtrB and the OMC decahaem c-Cyts MtrC and OmcA (Shi, et al., 2009). All these proteins
together form a pathway to transfer electrons from the quinone/quinol pool in the inner
membrane to the periplasm and then to the outer membrane where MtrC and OmcA can either
transfer electrons directly to the surface of electrode materials or to soluble redox mediators.
Although several studies have investigated the behavior of isolated OMCs attached on
electrodes (Eggleston, et al., 2008, Firer-Sherwood, et al., 2008b, Hartshorne, et al., 2007,
Hartshorne, et al., 2009, Meitl, et al., 2009, Pankhurst, et al., 2006, Turner, et al., 1999), or
embedded in microbial biofilms of Shewanella (Carmona-Martínez, et al., 2011, Coursolle, et
al., 2010, Liu, et al., 2011, Meitl, et al., 2009, Nakamura, et al., 2009a, Okamoto, et al., 2011,
Okamoto, et al., 2009), the role of these cytochromes in the heterogeneous ET across the
biofilm/electrode interface is far from clearly understood.
Surface-enhanced resonance Raman scattering (SERRS) is a sensitive and selective technique
able to probe surface-confined heme species in the sub-monolayer coverage (Khoa Ly, et al.,
2011). This approach, often used to study isolated proteins immobilized on nanostructured (i.e.
surface-enhanced Raman active) electrodes, is also applicable to multiheme proteins embedded
within intact electrochemical-active microbial biofilms (Millo, et al., 2011). In this case, SERRS
probes selectively the heme groups of the OMCs in the close vicinity of the electrode surface,
revealing important structural information, such as the oxidation-, the coordination-, the spin-
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state, and the nature of the axial ligands of the central iron atom of these proteins. When
performed in combination with electrochemical techniques, stationary and time-resolved
SERRS reveal unprecedented insights into the thermodynamics and the kinetics of the
heterogeneous electron transfer across the bacteria/electrode interface.
The aim of this study is to achieve a better understanding of the ET in microbial biofilms of S.
putrefaciens NCTC 10695. By measuring the electrochemical and spectroscopic properties of
microbial cells embedded in their natural biofilm habitat, a more realistic picture on the natural
electron transfer will be provided.
4.2 Materials and methods
4.2.1 Materials and methods
All chemicals were of analytical or biochemical grade and were purchased from Sigma-Aldrich
and Merck. If not stated otherwise, all potentials provided in this article refer to the Ag/AgCl
reference electrode (sat. KCl, 0.195 V vs. SHE). All microbial experiments were performed
under strictly sterile conditions.
4.2.2 Cell cultures and media
Sterilized LB broth (Ringeisen, et al., 2006), and minimal media (Bretschger, et al., 2007) (18
mmol/ L Sodium lactate, PIPES buffer 15.1 g×L-1
; NaOH 3.0 g×L-1
; NH4Cl 1.5 g×L-1
; KCl 0.1
g×L-1
; NaH2PO4∙H2O 0.6 g×L-1
; NaCl 5.8 g×L-1
; Wolfe’s mineral solution 10 mL×L-1
(Atlas,
1993); Wolfe’s vitamin solution 10 mL×L-1
(Atlas, 1993); Amino acid solution 10 mL×L-1
(Bretschger, et al., 2007)) for liquid cultures and LB/agar (Merck KGaA, Germany) plates were
used for culture maintenance.
Single colonies on LB/agar plates freshly streaked from a frozen glycerol stock culture
(Coursolle, et al., 2010) of S. putrefaciens wild-type NCTC 10695 (Nealson and Myers, 1992,
Pivnick, 1955, Venkateswaran, et al., 1999, Vogel, et al., 1997) (DSM No.: 1818, DSMZ -
German Collection of Microorganisms and Cell Cultures GmbH, Braunschweig, Germany);
were transferred to 15 mL of LB broth and incubated aerobically at room temperature while
shaking at 100 rpm (Universal shaker SM 30 A, Edmund Bühler GmbH, Germany) for 48 h
(Carmona-Martínez, et al., 2011). Afterwards 15 mL of culture were spun down at 3000 rpm
during 10 min (Heraeus Megafuge 1.0, Germany). The pellet was resuspended in 15 mL of
minimal media and transferred to a sterile Erlenmeyer flask with 185 mL of minimal media for
72 h of cultivation using the same conditions. Finally 200 mL of media were centrifugated at
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3000 rpm during 10 min, the pellet was resuspended in 15 mL of minimal media and injected in
the electrochemical cell.
4.2.3 Electrochemical set-up for the growth of anodic electrocatalytic biofilms
Sealed vessels (200 mL) served as the electrochemical cell hosting the three-electrode
arrangement (Fig. 4-2). The bioelectrochemical experiments were conducted under strictly
sterile conditions and under potentiostatic control (Potentiostat/Galvanostat VMP3, BioLogic
Science Instruments, France) utilizing a three-electrode arrangement, with a silver ring working
electrode (0.7 cm2) electrochemically roughened according to the procedure described elsewhere
(Wackerbarth, et al., 1999), a polycrystalline carbon rod counter electrode (CP Graphite GmbH,
Germany, 7.0 cm height x 1.0 cm diameter) and Ag/AgCl reference electrode (sat. KCl, 0.195 V
vs. SHE, Sensortechnik Meinsberg, Germany). The carbon electrode was glued with stainless
steel wire (AISI 304, Fe/Cr18/Ni10, Goodfellow GmbH, Nauheim, Germany) using a two-
component resin (Epoxy resin HT 2 + Hardener HT 2, HY-POXY® Systems, SC, USA) mixed
with carbon black particles (Vulcan XC-72, Cabot Corporation GMBH, Frankfurt am Main,
Germany).
Figure 4-2 Electrochemical half cell set-up under potentiostatic control. Insert shows a
photograph of the nanostructured silver ring working electrode.
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4.2.4 Growth of anodic electrocatalytic biofilms
The biofilm growth was performed in semi-batch chronoamperametric experiments at +0.05 V
at 30 °C (Owen, et al., 1978) with a regular Shewanella cells addition procedure (Baron, et al.,
2009) and media replacement as described below. For these potentiostatic biofilm growth
experiments, initial aerobic conditions were assured according to previous reports (Biffinger, et
al., 2009). Therefore, before adding the growth media into the electrochemical cell (Rosenbaum,
et al., 2010a, von Canstein, et al., 2008) filtered air was pumped into the fresh minimal media
using a fish pump (Elite air pump 799, Rolf C. Hagen Corp., MA, USA) per 200 mL for more
than 1 h. Fresh electron donor and nutrients were supplied about every 72 h by removing 160
mL of culture (representing 80% of the cell volume) and replacing with fresh minimal media
and fresh Shewanella cells according to Section 4.2.2.
4.2.5 Cyclic voltammetry
Cyclic voltammograms were recorded for non-turnover conditions according to (Carmona-
Martínez, et al., 2011) . Potentials were applied from -500 to +50 mV (vs. Ag/AgCl) at a scan
rate of 1 mV s-1
with continuous monitoring of the current response.
4.2.6 Electrochemical data processing
All data are based on experiments of 3 independent biofilm replicates and the standard
deviations are presented. The maximum current density, jmax, (calculated per projected electrode
surface area) was taken as a measure of activity and was analyzed during at least 2 semi-batch
cycles for established microbial biofilms. For in-depth data analyses of the cyclic
voltammograms the open–source software SOAS (Fourmond, et al., 2009) was used for baseline
(capacitive current) correction for non-turnover conditions.
4.2.7 Spectroelectrochemical set-up for SERRS measurements
Electrochemical experiments were carried out in a homemade spectroelectrochemical cell
working in the three electrode configuration and controlled by a μAutolab potentiostat (Eco
Chemie, Utrecht, The Netherlands). The three electrodes were a biofilm-coated (vide supra) Ag
ring electrochemically roughened (see 4.2.3), a Pt coil, and a Ag/AgCl (3.0 M KCl) (Dri-Ref,
WPI Berlin, Germany), acting as working, counter, and reference electrode, respectively.
4.2.8 SERRS measurements
SERRS spectra were obtained using a confocal Raman spectrometer (LabRam HR-800, Jobin
Yvon) with a spectral resolution of 2 cm-1
and an increment per data point of 0.57 cm-1
. SERR
spectra were obtained with 413 nm excitation (Coherent Innova 300 c Krypton cw-laser) with a
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laser power of 1.0 mW on the sample. The laser beam was focussed by a Nikon 20x objective
with a working distance of 20.5 mm and a numeric aperture of 0.35. Further details of the
experimental set-up are given elsewhere (Millo, et al., 2011).
4.3 Results and discussion
4.3.1 Bioelectrochemical current production
The biofilms were grown at a constant potential on roughened (i.e. SER-active) silver electrodes
using 18 mM sodium lactate as substrate (see above for experimental details). These biofilms
produced a maximum chronoamperometric current density of 3.03 ± 0. 13 μA cm-2
(Fig. 4-3),
which is in good agreement with previous studies using aerobe graphite anodes (see chapter 2
and 3).
Figure 4-3 Chronoamperometric curve of a biofilm formation using a silver ring electrode
poised at +0.05 V in a batch experiment using 18 mM sodium lactate as the substrate and S.
putrefaciens cells as biocatalyst.
The voltammetric behavior of the biofilms was monitored under non-turnover conditions (i.e.,
without sodium lactate). Fig. 4-4 shows the CV behavior of such a biofilm for non-turnover
conditions. The redox couple that is proposed to be involved in the DET (i.e. the one ascribed to
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the oxidation/reduction of the heme groups of the surface-confined OMC), Ef,1 is centered at a
formal potential of -233 mV (vs. the Ag/AgCl reference electrode). The main overall shape and
peak positions of the cyclic voltammogram shown in Fig. 4-4 are very similar to those obtained
on graphite electrodes in previous studies (see chapter 3), showing that biofilm formation is not
affected by the nature of the electrode material. Furthermore, this finding reveals that the
nanostructured silver electrode and specifically the inevitable traces of AgI cations do not
provide a toxic environment for Shewanella putrefaciens, confirming recent observations by
Preciado-Flores and co-workers (Preciado-Flores, et al., 2011) who were able to grow
Shewanella oneidensis cells in the presence of silver nanoparticles and nanowires.
Figure 4-4 A) CV of the active biofilm formed on a silver ring electrode under non-turnover
conditions (i.e. in the absence of the substrate sodium lactate) at a scan rate of 1 mV s-1
. B)
Respective SOAS baseline corrected curves.
Figure 4-5 shows the SERR spectra of a microbial biofilm grown on a roughened Ag electrode
and measured at different applied potentials in the absence of metabolic substrate. Spectral
analysis allows ascribing the SERRS signal to one or more heme species in the vicinity of the
electrode surface. Admittedly, the fingerprint region differs from that of a c-type heme, lacking
the typical band centered at 418 nm ascribed to the CCC vibrational mode of the heme (not
shown). Therefore, the type of heme cannot be unambiguously assigned. Moreover, the
frequency of the oxidation marker band 4 (in Fig. 4-5 at 1374 nm) has been found to vary
significantly. Although the high frequency values clearly show that the heme is oxidized, the
origin of the frequencies scattering obtained for different samples (1370-1380 cm-1
) and for the
same sample measured on two different days (1375-1380 cm-1
) is still unknown. Remarkably,
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no correlation with the sample treatment and/or preparation has been observed. Interestingly,
this frequencies scattering has not been observed for the other vibrational modes (i.e the 3, 2,
and 10). These frequencies are in agreement with a six-coordinated low-spin Fe atom with two
His residues acting as axial ligands (Oellerich, et al., 2002).
Figure 4-5 SERR spectra of the reduced (upper spectrum) and oxidized (lower spectrum)
OMCs, obtained at -425 and 0 mV, respectively. The spectra were obtained with excitation at λ
= 413 nm, laser power of 1 mW, and an acquisition time of 90 s. Potentials refer to the Ag/AgCl
(KCl 3 M) reference electrode (210 mV vs. SHE).
As shown in Figure 4-5, spectra recorded at two different poised potentials are very similar.
Remarkably, the spectrum at -475 mV does not display the expected band downshift of the 4
upon electrochemical reduction. This finding is indicative that the heme species detected by
SERRS – probably the heme groups of surface-confined OMCs – do not promote the electron
transfer. OMC reduction has been attempted by applying a negative potential in the presence of
(i) O2, (ii) the soluble redox mediator riboflavin, (iii) O2 and riboflavin together. In all cases no
reduction has been observed. Full OMC reduction has been achieved by exposing the biofilm at
open circuit potential to the soluble reducing agent dithionite (Na2S2O4). This procedure has
lead to the immediate OMC reduction, as proven by the downshift of the 4 to 1360 cm-1
(not
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shown). However, after this harsh chemical treatment, the biofilm lost its capability of
generating electrical current. In fact, chronoamperometric experiments performed in the
presence of lactate did not show the recovery of catalytic activity.
4.4 Conclusions
The experiments here presented, performed on microbial biofilms of S. putrefaciens NCTC
10695, have shown that OMCs do not contribute to the heterogeneous ET across
bacteria/electrode interface. These studies have been performed on biofilms grown on
nanostructured Ag electrodes at the poised potential of +50 mV (vs. Ag/AgCl). Although these
conditions allow the formation of a biofilm on the Ag electrode, they may have a negative
impact on the amount of OMCs expressed by the bacteria (see chapter 3). In fact, optimal
biofilm growth requires more positive potentials. However, these conditions cannot be met by
the Ag substrate, which undergoes oxidation at potentials higher than +150 mV (vs. Ag/AgCl).
The development of novel analytical strategies to overcome this limitation is presently under
evaluation in our groups. Furthermore, under the supervision of Dr. Millo, I will perform further
experiments at the Vrije Universiteit Amsterdam that will allow us achieve a better
understanding of the electron transfer in microbial biofilms of S. putrefaciens.
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CHAPTER V CHAPTER V
5 Electrospun and solution blown three-dimensional
carbon fiber nonwovens for application as electrodes in
microbial fuel cells
5.1 Introduction
There is an ever increasing interest in the research community and industries in electrospinning
and solution blowing of microfiber, nanofiber and nonwovens for applications such as
biomedical (Agarwal, et al., 2008, Reneker, et al., 2007), tissue engineering (Pham, et al., 2006,
Wise, et al., 2009), drug release (Gandhi, et al., 2009, Srikar, et al., 2008, Wang, et al., 2010),
agriculture (Hellmann, et al., 2011), etc. Recently high-temperature electrospun nonwovens
were used as separators in lithium-ion batteries (Bansal, et al., 2008). Typical electrospun
nonwovens represent themselves thin, practically two-dimensional, sheets of low permeability.
For a number of applications, e.g. tissue engineering, three-dimensional, fluffy, nonwovens
could be of great interest.
Preparation of three-dimensional nanofiber nonwovens by remodeling of nanofiber yarn
electrospun onto water surface was recently reported in biomedical context (Teo, et al., 2008).
Three-dimensional multi-layered cell–nanofiber constructs for tissue engineering were also
prepared by a kind of layer-by-layer electrospinning (Yang, et al., 2009). The new capabilities
of the preparation techniques and versatility of the resulting nonwoven materials raise the
question if bioelectrochemical systems (BES) like microbial fuel cells can benefit from these
developments and their performance can be increased further more. The great majority of
current microbial BES utilizes electroactive microbial biofilms as electrocatalyst to facilitate the
anodic substrate oxidation.
Key players in such biofilms are electroactive bacteria like Geobacter or Shewanella species
that are often also referred to as electricigens or exoelectrogens (Gorby, et al., 2006, Logan,
2009, Lovley, 2006, Rabaey, et al., 2010, Rabaey, et al., 2007). Exploiting electroactive bacteria
in BES, e.g. in microbial fuel cells or electrolysis cells (Rabaey, et al., 2010), promises a great
potential in the context of sustainable energy supply and handling. A major challenge in this
context is an increase in the performance of such systems. During the past decade the average
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current densities of biofilm anodes have already increased significantly from milliampere per
square metre level to between 7 and 10 A m-2
. This increase can be mainly attributed to the
improvement of the microbial biofilms, e.g. via sophisticated enrichment and acclimatization
procedures (Kim, et al., 2004, Liu, et al., 2008, Rabaey, et al., 2004). Recent experimental
results and theoretical considerations suggest that a further improvement above 10 A m-2
may be
hampered by limitations imposed by kinetics associated with electron and proton transfer as well
as substrate diffusion within the biofilm (Torres, et al., 2010, Torres, et al., 2008). Since the
improvement of the biological component becomes increasingly difficult, the improvement or
even the tailoring of the electrode materials becomes an important task. In the past, such
electrode tailoring has already been shown to be successful for microbial fuel cell based on
suspended bacterial cultures (Rosenbaum, et al., 2006, Schröder, et al., 2003). One strategy to
enhance the performance of biofilm anodes is the improvement of the electrode surface
properties by surface treatment procedures such as ammonia treatment, polymer modification or
surface oxidation (Cheng and Logan, 2007, Scott, et al., 2007, Wang, et al., 2009). These
measures are likely to improve biofilm–electrode interactions and thus the rate of electron
transfer. A further, promising path is the increase of the active surface area, for example by
means of brush (Logan, et al., 2007) or fiber electrodes e.g. ref. (Liu, et al., 2010c), and 3D
electrode materials e.g. ref. (Zhao, et al., 2010b). This path already delivers promising results as,
for example in ref. (Zhao, et al., 2010b), which combines a 3D electrode structure and a
conductive polymer as an immobilized mediator. The experiments of ref. (Zhao, et al., 2010b)
yielded the highest reported current density of a biofilm electrode (about 24 A m-2
). The present
work aims to exploit high surface area electrospun and solution-blown carbonized nonwovens to
further enhance the current density of BESs anodes. Three different fiber mat materials
(nonwovens) were studied in comparison to conventional polycrystalline graphite and carbon
felt, i.e.: i) 3D porous carbon fibers, produced by gas-assisted electrospinning (hereafter denoted
as GES-CFM); ii) Electrospun carbon fibers (ES-CFM); and iii) Solution-blown carbon fibers
(SB-CFM). The latter two materials were additionally modified to incorporate 15% carbon black
(CB) to increase porosity and conductivity of the resulting nonwovens. These modified
materials are denoted as ES-CFM15%CB and SB-CFM15%CB. In order to study and compare
the bioelectrocatalytic substrate oxidation at all above-mentioned electrode materials,
preselected wastewater-derived mixed culture biofilms were grown under potentiostatic control.
The bioelectrocatalytic performance was then analyzed in semi-batch, chronoamperometric
experiments by measuring the projected (geometric) current density at the respective materials.
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5.2 Materials and methods
5.2.1 Carbon fiber preparation
Three techniques for the preparation of three-dimensional carbon fiber mats (CFM) were
employed for this study: electrospinning (see Fig. 5-1A, samples labeled ES), solution blowing
(SB) and gas-assisted electrospinning (GES).
Figure 5-1 (A) Schematic drawing of an electrospinning setup (derived from ref. (Greiner and
Wendorff, 2007)). Solution blowing differs from electrospinning by the use of a high-speed
nitrogen jet flow (230–250 m s-1
) instead of a high voltage electric field to accelerate and stretch
the polymer solution into a fibrous form (Sinha-Ray, et al., 2010). (B) Electrochemical cell for
the simultaneous study of different electrode materials.
5.2.1.1 Gas-assisted electrospinning carbon fiber mat (GES-CFM)
A solution of poly(acrylonitrile-co-itaconic acid-co-butyl acrylate) (monomer ratio AN/IA/BA =
46/3/1; [η]25°C = 2.4 dL g-1
measured in DMAc) 11 weight% in DMSO/DMF/acetone (16/15/5)
was used for air-assisted electrospinning. The electrode distance was 100 cm between the
electrodes at a voltage of 40 kV m-1
. The carbonization process was performed in a high-
temperature furnace using the following protocol:
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i) Heating up to 230°C at a rate of 20°C min-1
in air and annealing for 2 h for stabilization of the
precursor ultrafine fibers; ii) Heating up to 350°C at a rate of 5–10°C min-1
in nitrogen
atmosphere and annealing at 350°C for 20 min; iii) Heating up to 750°C at a rate of 3–5°C min-1
in nitrogen atmosphere and annealing at 750°C for 20 min; and iv) Heating up to 1000°C at a
rate of 3–5°C min-1
in nitrogen atmosphere and annealing at 1000°C for 1 h to complete the
carbonization process.
5.2.1.2 Electrospun carbon fiber mat (ES-CFM)
12% polyacrylnitrile (PAN) solutions containing 5, 7, 9, 11 and 15 wt% of carbon black (CB)
nanoparticles were electrospun. The resulting fiber mats had a very loose three-dimensional
structure with high porosity (with pores approximately from ten to hundreds of micrometres in
comparison with the ordinary electrospun fiber mats with pores of about 3 to 5 mm). The
resulting PAN fiber mats were placed in a furnace and stabilized in air for 20 min at 270°C, then
carbonized in nitrogen at 1100°C (the ramp rate was 4°C min-1
between the room temperature,
280°C, and 1100°C plateaus).
5.2.1.3 Solution-blown carbon fiber mat (SB-CFM)
Blends of PAN solutions with CB were solution blown as described in ref. (Sinha-Ray, et al.,
2010). The blends were prepared using the following steps. Initially a 12% PAN solution in
DMF was prepared. Then, 1 g of 12 wt% PAN solution, 0.12 g of CB and 4 g of DMF in a 20
mL vial were sonicated for 45 min. During sonication CB clusters were broken, dispersed and
coated by PAN, which prevented coalescence of particulates. Then, 4.32 g of DMF and 0.68 g
of PAN were added to the vial and kept on a hotplate overnight under permanent stirring.
In this way a blend of 8 wt% PAN solution with 15 wt% CB was prepared and solution blown
(Sinha-Ray, et al., 2010). The carbonization was achieved by heating of the PAN fibers in open
air at 350°C for 3 h and then in nitrogen atmosphere at 1050°C for 1 h.
Porosity data are provided as a fraction of the void volume in the total volume:
total
solid
total
void
V
V
V
V 1 Equation 5-1
Here, Vvoid is the volume of pores, Vsolid is volume of solid, Vtotal is the total or bulk volume of
material including the solid and pore volume. The volume of the fiber material can be expressed
as V = Aδ, with A being the geometric surface area and δ the equivalent thickness of the fully
compressed fiber mat.
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Using a constant geometric surface area, Equation 5-1 can be written as Θ = 1 - δsolid/δtotal. Here,
δtotal represents the effective thickness of the porous fiber mat, δsolid is the equivalent thickness of
the fully compressed solid material calculated using the mass, m, of the fiber mat and the density
of graphite, ρ, as δsolid = m/ρA.
5.2.2 Electrode preparation
All above-mentioned materials were utilized as electrodes for the growth and investigation of
(anodic) electrocatalytic biofilms. For electrode preparation, either the electrode materials
prepared in this work or commercial carbon felt (Weichfilz, SIGRATHERM, SGL Carbon
GmbH, Meiningen, Germany). They were cut into 1x2 cm2 pieces and glued onto graphite foil
paddles (serving as an inert electrode backbone material; Chempur©, Karlsruhe, Germany)
using a two-component resin mixed with carbon black particles (Vulcan XC-72). Polycrystalline
carbon was used as carbon rods (CP Graphite GmbH, Germany).
5.2.3 Bioelectrochemical experiments
All bioelectrochemical experiments were conducted under strictly anoxic conditions and under
potentiostatic control. The potentials provided in this manuscript refer to the Ag/AgCl reference
electrode (sat. KCl, 0.195 V vs. SHE, Sensortechnik Meinsberg, Germany). All microbial
biofilms were grown in potentiostatic half-cell experiments at 0.2 V at 35°C (Patil, et al., 2010).
To assure comparability and reproducibility up to six electrodes were measured simultaneously
in one electrochemical cell (cf. Fig. 5-1B).
For this purpose, an Autolab 30 potentiostat (Ecochemie, the Netherlands) equipped with six
array channels was employed. The bacterial growth medium was prepared as reported in ref.
(Liu, et al., 2008) with acetate serving as a substrate of the medium (pH 6.8). The bacterial
source for the primary biofilm formation was wastewater from the wastewater treatment plant
Steinhof, Braunschweig.
The reported current densities refer to secondary, i.e. preselected, biofilms (gained by a
procedure as described in ref. (Liu, et al., 2008). They are based on three independent biofilm
replicates with at least two feeding cycles per biofilm, exhibiting a reproducible maximum
current density. Cyclic voltammograms were recorded for turnover and non-turnover conditions
according to ref. (Fricke, et al., 2008).
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5.3 Results and discussion
5.3.1 Biocatalytic current generation at modified carbon electrodes
Table 5-1 summarizes the projected (geometric) current density data and physical parameters of
the tested electrode materials. The current density data correspond to the maxima of the
respective semi-batch experiment (Fig. 5-2), averaged over at least three independent
experimental sets. Table 5-1 illustrates that conventional polycrystalline carbon graphite shows
the lowest bioelectrocatalytic current density of all tested samples. The corresponding current
value of 13 A m-2
is still quite high. This is explained by the fact that in the present work we use
preselected (secondary) biofilms with an optimized performance (Liu, et al., 2008), in
combination with the reaction temperature of 35°C (Patil, et al., 2010).
Table 5-1 Cumulative data on electrocatalytic current densities obtained at different electrode
materials. The substrate was 10 mM Sodium acetate.
Material Current densitya /
A m-2
Specific weight /
g m-2
Specific current
density/ mA g-1
Polycrystalline graphite 13 NA NA
Carbon Felt 16 333 48
GES-CFM 30 42 714
ES-CFM 21 126 243
ES-CFM15%CB 15 88 172
SB-CFM 17 437 56
SB-CFM15%CB 21 183 128
aProjected (geometric) current density, NA—not applicable.
Carbon felt revealed a higher (by 23%) performance than polycrystalline graphite. The highest
current density in this study was obtained with GES-CFM. As illustrated in Fig. 5-2, biofilms
grown on GESCFM-electrode delivered maximum current densities of 30 A m-2
. To the best of
our knowledge these are the highest current densities so far achieved with an electroactive
microbial biofilm.
The other material types, ES-CFM and SB-CFM, also yielded current densities higher than
commercial carbon felt, and up to 21 A m-2
(Table 5-1). Interestingly, the addition of carbon
black so far did not deliver unambiguous results. In the case of the solution blown material, SB-
CFM, addition of 15% CB increased the current density from 17 A m-2
to 21 A m-2
.
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On the other hand, the electrospun ES-CFM did not benefit from adding carbon black. In the
latter case the current density decreased from 21 to 15 A m-2
. The reason for this behavior is not
known so far and will require further investigation.
Figure 5-2 Biocatalytic current generation at a GES-CFM modified carbon electrode in a model
semi-batch experiment. The GES-CFM electrode was modified by a wastewater-derived
secondary biofilm grown in a half-cell experiment under potentiostatic control. The electrode
potential was 0.2 V.
The excellent performance of GES-CFM, the porous carbon mat produced by gas-assisted
electrospinning, is realized at extremely low specific weight (see Table 5-1). In comparison to
commercial carbon felt (specific weight 333 g m-2
) that of GESCFM was decreased by 87%
(specific weight 42 g m-2
). GES-CFM further possesses an extremely low effective density of
about 18 kg m-3
-which is an order of magnitude lower than that of conventional carbon-based
materials, such as carbon felt (about 100–180 kg m-3
).
5.3.2 Analysis of electroactive biofilms grown at modified carbon electrodes with
Scanning electron microscopy
The high porosity of the material is achieved by enhanced electric fiber–fiber repulsion and
mechanical restrictions when layers of fibers are deposited one by one. Further, the presence of
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a gas flow disturbs the stable electrospinning jet and causes the fiber to form loose fiber
aggregates. Due to the high solvent content during the preparation these fiber aggregates form
interconnections (Fig. 5-3E), which lead to a high mechanical stability as well as a high
electronic conductivity of the fiber mats.
Figure 5-3 Scanning electron microscopic images of (A) carbon felt, (B) an electroactive
biofilm grown at carbon felt, (C) GES-CFM, (D) an electroactive biofilm grown at GES-CFM,
(E) high resolution image of GESCFM illustrating the occurrence of inter-fibre junctions, and
(F) crosssectional view of GES-CFM electrode.
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Thus, the excellent bioelectrocatalytic performance of this material can be attributed to a
structure that provides a habitat for the growth of electroactive bacteria up to a maximum
density supplemented by efficient substrate supply through the open pore structure. The
interconnections between the individual fibers of the nonwoven allow the formation of cross-
linked three-dimensional biofilms (Fig. 5-3D) that benefit from an optimum electron transfer
and conduction.
The origin of the performance differences between the different electrospun and solution blown
materials is so far not fully understood. All materials possess an extremely high porosity of up
to 99%. Porosity represents the fraction of void space in the material and is of utmost
importance. Indeed, high porosity not only lowers the amount of material to a minimum but also
maximizes penetration of microorganisms and the diffusional substrate supply. However,
parameters like the occurrence of cross-linking points (cf. Fig. 5-3E) may represent a key factor
that lead to an additional performance gain.
5.3.3 Cyclic voltammetry of electroactive biofilms grown at modified carbon electrodes
Cyclic voltammograms were recorded under turnover as well as non-turnover conditions in
order to study a potential effect of the electrode material on the mechanism and thermodynamics
of the biocatalytic substrate conversion. Fig. 5-4 depicts cyclic voltammograms of the
electroactive biofilms at GES-CFM under turnover as well as non-turnover conditions. The
results are typical for wastewater-derived electroactive biofilms and biofilms of Geobacter
sulfurreducens.
Thus, the cyclic voltammograms of the biofilms under non-turnover conditions reveal redox
systems with an average formal potential of -329 mV, which is in accordance with the literature
data. In particular, in previous studies, formal potentials of -324 mV were achieved for
secondary mixed culture biofilms (Liu, et al., 2008) and -331 mV for biofilms of G.
sulfurreducens (Fricke, et al., 2008), both grown at polycrystalline graphite.
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In light of the recent experimental proof that G. sulfurreducens represent a strongly dominating
species in acetate-grown wastewater-derived biofilms (Harnisch, et al., 2011), the present
voltammetric results confirm that the thermodynamics of bioelectrocatalytic energy conversion
is not determined or affected by the fiber electrode material but is fully governed by the
extracellular microbial electron transfer (e.g., by the redox potential of the outer membrane
cytochromes).
This is an important issue since any deterioration of the thermodynamics by the introduction of
additional redox cascades, e.g., via the use of immobilized redox mediators, would lower the
electrochemical energy gain and thus the energy efficiency of a potential bioelectrochemical
device.
Figure 5-4 Cyclic voltammograms of an electroactive biofilm grown at GESCFM. The
voltammograms were recorded under turnover conditions [in the presence of substrate (10 mM
acetate), curve A], as well as nonturnover conditions (the absence of substrate, curve B). The
biofilm was a wastewater-derived secondary biofilm grown at a potential of 0.2 V under
potentiostatic control. The scan rate was 1 mV s-1
.
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5.4 Conclusions
In conclusion, it has been shown that three-dimensional electrospun and solution-blown carbon
fiber nonwovens represent highly interesting and promising electrode materials for microbial
fuel cell applications. They combine the use of a minimum amount of carbon and great
performance. Thus, current densities of up to 30 A m-2
were achieved, which represent the
highest values demonstrated at electroactive biofilms to date. These current densities were
achieved without chemical surface modification, which may represent an advantage with respect
to longevity.
Based on this initial study, further and systematic investigations are required to fully exploit the
potential of this class of materials. A special emphasis of further investigations must be on tests
designed to elucidate the long-term behavior of the new electrode materials and the resistivity
against clogging of the pore structures. Further, a theoretical model is under investigation to
correlate and predict structure–property relationships of this new class of BES anode materials.
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CHAPTER VI CHAPTER VI
6 Electrospun carbon fiber mat with layered architecture
for anode in microbial fuel cells
6.1 Introduction
Bioelectrochemical systems like Microbial fuel cells (MFCs), considered to be a green energy
conversion technology, have recently attracted appreciable attention for the conversion of
chemical energy to electricity (Logan, 2009, Logan and Regan, 2006b, Rabaey and Verstraete,
2005). The creative idea of MFCs originated from the discovery that microbial biofilms can be
adapted to extracellular electron transfer (Rabaey, et al., 2010). Numerous studies have found
that the key players in these biofilms are electroactive bacteria (Logan, 2009, Logan and Regan,
2006a), like members of the Geobacter (Harnisch, et al., 2011, Lovley, 2006) or Shewanella
(Gorby, et al., 2006) families.
The performance of MFCs, to a large extent, depends on the density of electrochemically active
biofilms on the anode. Most of the literature reported the use of two-dimensional MFC anodes
(Adachi, et al., 2008, Qiao, et al., 2008, Qiao, et al., 2007). The 2D nature of anode allowed
biofilms to grow only in one direction and made a layer on the surface. Some new materials,
such as carbon nanotube/polyaniline composite (Qiao, et al., 2007), Titanium
dioxide/polyaniline composite (Qiao, et al., 2007), and mediator mobilized carbon (Adachi, et
al., 2008) have been used for anode in MFCs and generated exciting current density.
Thus, e.g. in ref. (Adachi, et al., 2008) a current density up to 1.2 mA cm-2
has been generated
by using 9,10-anthraquinone-2,6-disulfate modified graphite felt. Carbon materials with three-
dimensional porous architecture have also been used for anodes in MFCs to increase active
surface for biofilm growth, like reticulated vitreous carbon (He, et al., 2005), graphite fiber felt
(Chaudhuri and Lovley, 2003, Liu, et al., 2010c, Zhao, et al., 2010b) and brush materials
(Logan, et al., 2007) and very promising results have been obtained. Especially for large scale
applications, e.g. wastewater treatment and the accompanying MFC up-scaling, the increase of
the overall MFC performance is still a big task.
Electrospinning is a unique process that effectively produces small fibers with diameter ranging
from a few nanometers to several microns (Greiner and Wendorff, 2007). Electrospun fiber mats
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with advantages of small diameter and high specific surface area have been widely used in many
fields, which were highlighted in many review articles (Greiner and Wendorff, 2007, Thavasi, et
al., 2008). Carbon nanofiber mats (CFMs) obtained by carbonization of electrospun
polyacrylonitrile (PAN), showed a high porosity, e.g. >85%. Because of their high surface area
and porosity, the carbonized electrospun fiber mats (Hou and Reneker, 2004, Wang, et al.,
2002b, Zhou, et al., 2009, Zussman, et al., 2005) have been used as electrodes in
electrochemical cells (Guo, et al., 2009, Ji and Zhang, 2009, Kim, et al., 2007, Kim and Yang,
2003, Kim, et al., 2006). Electrospun CFMs (ECFMs) could also be used as high performance
anode in microbial fuel cells. In Chapter 5, three-dimensional ECFM prepared by gas-assisted
electrospinning, has been proved to be an efficient anode in MFCs and generated the highest
anodic current density of 3.0 mA cm-2
known - to the best of my knowledge - till date (Chen, et
al., 2011). This high anodic current density has been believed to be attributed to the large
surface area and ultrahigh porosity for growth of high density biofilm.
In this Chapter, the concept of continuous, layered anode biofilms for microbial
bioelectrochemical systems is introduced. Three-dimensional CFMs with layered architecture
(hereafter denoted as layered-CFMs) were prepared to grow continuous, layered electroactive
biofilms with high density. Natural cellulose paper (NCP) was used as support for layer-by-layer
electrospinning (LBL-electrospinning) of PAN fiber layers. The layered-CFM has an
architecture of alternating cellulose-based and electrospun PAN-based carbon fiber layers, and
shows an ultrahigh porosity of 98.5%. The layered-CFM was used as MFC anode and the
biofilm growth characteristics were investigated.
6.2 Materials and methods
6.2.1 Carbon fiber preparation
10 wt% PAN (Mw=210 k) solution in dimethylformamide (DMF) was prepared for
electrospinning. The typical conditions used for electrospinning were electric field of the order
of 60 kV m−1
, voltage of 10 kV, and the distance between two electrodes was 20 cm. The
feeding rate of solution was 0.1 mL min−1
. Layered NCP-PAN fiber mat of ten layers (numbers
of PAN fiber layers) was prepared by layer-by-layer electrospinning of PAN fibers onto thin
NCP (in the form of tissue paper). Each PAN fiber layer was electrospun for about 5 min. All
fiber mat samples were dried in vacuum at 60 °C for subsequent carbonization.
The electrospun fiber mats and NCP were sandwiched between two graphite plates and
carbonized in a tubular stainless steel reactor by using the following protocol: 1) in air
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atmosphere, heating up to 230°C at a rate of 2°C min−1
and annealing for 3 h to finish the
stabilization process; 2) in N2 atmosphere, heating up to 500°C at the rate of 2°C min−1
and
annealing for 1 h, then heating up to 1000°C at a rate of 5°C min−1
and annealing for 1 h.
CFMs from NCP (CFM-NCP), two-dimensional typically electrospun PAN fiber mat (2D-
ECFM) and layered NCP-PAN (layered-CFMs) were prepared for subsequent measurement.
6.2.2 Electrode preparation
All above CFM electrodes and including commercial carbon felt (CCF) (SGL Carbon GmbH)
were utilized as electrodes for the growth and investigation of (anodic) electrocatalytic biofilms.
They were cut into 1×2 cm2 pieces and were glued onto graphite foil paddles (serving as current
collector) using a conductive resin produced from two-component resin mixed with carbon
black particles.
6.2.3 Bioelectrochemical measurements
All electrochemical experiments were carried out as half-cell experiments under potentiostatic
control, using a three-electrode arrangement consisting of a working electrode, an Ag/AgCl
reference electrode (sat. KCl, 0.195 V vs. SHE) and a graphite plate counter electrode (size of
4×5 cm2). The experiments were conducted under control of a potentiostat (Autolab
PGSTAT30, equipped with six array channels) at 35°C (Patil, et al., 2010). All electrodes were
put in one chamber, a potential of 0.2 V (vs. Ag/AgCl) was applied on working electrode, and
the current was recorded in real time. To assure comparability and reproducibility up to six
different anode materials were measured simultaneously in one electrochemical cell.
The medium for bacterial growth was prepared with 10 mM Sodium acetate in 0.05 M sodium
phosphate buffer solution (pH 6.8). The bacterial communities were secondary biofilms which
were preselected from wastewater (wastewater treatment plant Steinhof, Braunschweig)
following procedures described in Ref. (Liu, et al., 2008). The electrochemical performance
tests were conducted when the biofilm activity reached stationary level.
6.2.4 SEM imaging
The fixation and drying of biofilm samples for morphology characterization were prepared as
follows: 1) fixed by 5 wt% glutaric aldehyde in 0.05 M phosphate buffer solution (pH=7.0); 2)
dehydrated in a graded series of aqueous ethanol solution; 3) then taken out and naturally dried
at room temperature. Scanning electronic microscopy (SEM) images were obtained from JSM-
7500F equipment under a voltage of 5 kV. The energy dispersive X-ray analysis (EDX) was
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conducted by a CamScan SEM (Model S2-80DV). The mean pore size of the CFM samples was
measured by a capillary flow porometer (PMI, CFT-1200AEXL) using dry up/wet up method.
The ohmic resistance of carbon fiber mats was measured by stand four-point method using a
Keithley 2000 multimeter at room temperature.
6.3 Results and discussion
6.3.1 Properties and performance of carbon fiber mat electrode materials
Generally, NCPs in the form of tissue papers are made of cellulosic fiber pulp and are used in
daily life as toilet papers, paper handkerchief etc. The NCPs used in this work, were porous and
had a mean pore size of about 23 μm with layer thickness of about 20 μm, as shown in SEM
images (Fig. 6-1A and 6-1B). The resulting NCP-CFM exhibited a low electrical resistivity of
about 7.3 Ω cm. The properties of different CFMs are summarized in Table 6-1.
Table 6-1 Properties and anodic performance of carbon fiber mats.
Sample Mean pore
size/ μm
Porosity/
%
Resistivity/
Ω cm
Geometric current
density/ mA cm-2
CCF 47.0 95.7 0.2 1.21
NCP-CFM 38.0 93.6 7.3 0.53
2D-ECFM 0.6 90.0 8.0 0.17
Layered-CFM 2.3 98.5 2.0 2.00
The EDX spectrum shown in Fig. 6-1C indicated that the NCP-CFM contained a very small
amount of normal metal elements, e.g. Na, Al, K and Ca, in the form of phosphate or sulfate
salts. These elements were expected to show no adverse effects on the growth of
microorganisms.
Taking these features of NCPs into consideration, in this work, thin layers of NCP were selected
as support for LBL-electrospinning of PAN fibers to fabricate layered-CFM. In this layered-
CFM, the PAN fibers of about 500 nm diameter were assembled on the NCP (10–30 μm) layers
(Fig. 6-1D and 6-1E). The layered-CFM had a high porosity of 98.5% owing to a large gap of
over 50 μm between layers and small electrospun fibers diameter (Fig. 6-1E). This layered-CFM
structure as seen by SEM proved the hypothesis and would provide habitat for growth of layered
microbial biofilms.
The anodic performance of this layered-CFM was tested in a batch half-cell MFC. The
bioelectrocatalytic current generation curves were recorded versus time. They originated from
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the oxidation of the substrate (acetate) catalyzed by microbial biofilms at the working electrode
according to the following equation:
eHCOOHCOOCHcatalysebioelectro
872 223 Equation 6-1
Figure 6-1 A) Top view and B) cross-sectional view SEM images of carbon mat from TP; C)
EDX spectra of NCP-based carbon fiber; D) top view and E) cross-sectional view SEM images
of layered-ECFM; F) cross-sectional view SEM image of 2D-ECFM.
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6.3.2 Biocatalytic current generation at carbon fiber mat electrode materials
The bioelectrocatalytic current generation curves of five cycles are shown in Fig. 6-2. The
layered-CFM anode generated a maximum geometric current density of over 2.0 mA cm−2
. This
is higher (about 65%) than that obtained from CCF of 1.21 mA cm−2
and NCP-CFM with ten
layers of about 0.53 mA cm−2
.
The anodic current density from layered-CFM was ten times higher than that from 2D-ECFM of
only 0.17 mA cm−2
. The possible reasons for the high anodic current density from layered-CFM
are 1) the large space between layers provided room for high density biofilms growth and makes
the substrate supply into the layer easy, and 2) electrospun fiber layer with smaller diameter
induces microorganisms to form thick and stable layered-biofilms.
Figure 6-2 Biocatalytic current generation curves of carbon fiber mats in a half-cell experiment
measured at room temperature. Arrows represent replacement of medium.
6.3.3 Analysis of electroactive biofilms grown at carbon fiber mat electrode materials
with Scanning electron microscopy
To confirm the anodic performance, biofilms in the CFMs were investigated by scanning
electron microscope (SEM) (JSM-7500F). Thick and continuous layered biofilms were grown in
the layered-CFM, as shown in Fig. 6-3A to Fig. 6-3C. The biofilms in the first layer had a
thickness of around 10 μm (Fig. 6-3C), and it became thinner in the inner layers due to substrate
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supply limitation. It revealed that the thickness of biofilms in the inner layers can be increased if
the gap between layers further increased for efficient nutrition transportation. Due to the big
pore size in the CCF resulted by big fiber diameter, the microorganisms were wrapped around
the big carbon fibers but did not form continuous layered-biofilm over the whole fiber mat (Fig.
6-3D and E).
Figure 6-3 SEM images of biofilms in: A-C belong to layered-CFM; D and E belong to
commercial carbon felt; and F belongs to 2D-ECFM.
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For the 2D-ECFM, as shown in Fig. 6-3F, only a very thin biofilm was grown on the surface of
2D-ECFM owing to the small pore size among fibers (Fig. 6-1F). The small pore size hindered
the microorganisms going inside and generated extremely low current density of only 0.17 mA
cm−2
. In summary, here it has been showed that small fiber diameter and proper pore size
combined with sufficient three dimensionality are essential features for the growth of thick and
continuous biofilms as well as for generation of high current densities.
6.4 Conclusions
Layered-CFM had been prepared and used for the anode in microbial fuel cells. These results
show that carbon materials with layered architecture and small fiber diameter are suitable for
layered biofilms growth with high cell density. Thick and continuous layered biofilms were
grown on this layered-CFM and generated high current density. This investigation also revealed
that if the gap between the layers within the layered-CFM can be further increased, thick layered
biofilms would grow in every layers of the entire layered-CFM and much larger current
densities would be obtained. The cellulose-based carbon fiber mat might provide a low cost and
highly efficient electrode for the anode in microbial fuel cells.
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CHAPTER VII CHAPTER VII
7 Electroactive mixed culture derived biofilms in microbial
bioelectrochemical systems: the role of pH on biofilm
formation, performance and composition
7.1 Introduction
Electrochemically active microbial biofilms not only play a key role in environmental oxidation
reduction cycles, e.g. (Nielsen, et al., 2010), but also in microbial bioelectrochemical systems
(BES) (Rabaey, et al., 2010). Within this seminal technology microbial biofilms are exploited
for anodic oxidation reactions (Logan, 2009, Lovley, 2008b, Schröder, 2007) as well as cathodic
reduction reactions, e.g. (Harnisch and Schröder, 2010). These latter reactions may range from
the oxygen reduction in microbial fuel cells (MFC) to the reductive production and/ or
upgrading of chemicals, e.g. H2, in microbial electrolysers.
Common to the majority of these BES applications is the biofilm at the anode that is responsible
for the microbially assisted oxidation of the substrate (i.e. wastewater constituents). Except for
pure culture studies, which are highly relevant concerning the investigation of fundamentals, the
anodic biofilm in BES are generally formed from natural bacterial sources, i.e. inoculums, like
wastewater. The wastewater derived biofilms exploited in the initial phase of BES research
often possessed an only minor bioelectrocatalytic activity (Kim, et al., 2001) and consequently
different enrichment procedures were presented leading to an increased anodic biofilm
performance, see e.g. (Kim, et al., 2005, Liu, et al., 2008, Rabaey, et al., 2004).
Up to now, the majority of BES studies using mixed culture biofilms were performed using
laboratory conditions tailored towards highest activity, i.e. metabolic turnover, and thus
maximum current production. However, as BES technology has to be integrated into wastewater
treatment technology lines (Rozendal, et al., 2008) it has to be taken into account that the
biofilms may face different, often suboptimal and quickly varying abiotic conditions during their
formation and operation. This is especially a challenge when wastewater is used as feed, since
its quality changes quickly due to the amount and kind of the various inflow sources. Recently,
it has been demonstrated on the example of the operation temperature (Patil, et al., 2010) that
the influence of external environmental conditions can be severe.
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Concerning the influence of the pH-value in the anodic compartment in BES, all recent studies
were restricted to a comparably narrow pH-window around pH neutral, e.g. (Biffinger, et al.,
2008, He, et al., 2008, Hong, et al., 2009, Jadhav and Ghangrekar, 2009, Liu, et al., 2005, Puig,
et al., 2010). Furthermore, acidophilic (Borole, et al., 2008) or alkalophilic (Liu, et al., 2010b)
microorganisms were exemplarily studied for a potential application of BES under extreme pH
conditions. Yet, as all these studies were performed in entire MFC devices, in which not only a
potential pH dependent biofilm performance contributes to the overall BES behaviour, but also
the pH-dependence of several technical operational parameters like that of the ion transfer
between anode and cathode (Harnisch and Schröder, 2009, Rozendal, et al., 2006a) or the
cathodic oxygen reduction reaction (Zhao, et al., 2006). Whereas the latter technological aspects
can be compensated by adequate technical measures (e.g. tailored geometries and materials), the
anodic biofilm may determine the overall pH-window of possible BES application. The aim of
this study is to provide information on the pH-influence of these biofilms from the short to
medium time frame (hours to days), as pH-associated metabolic adaptations take place within
minutes to hours (Siegumfeldt, et al., 2000).
Thus, in the present study the influence of the pH during formation and operation of natural
community derived anodic microbial biofilms was investigated using pH-values between pH 3
and pH 11. The biofilm formation, electrochemical performance, and redox-behaviour were
explored. Furthermore, the microbial structures and compositions of exemplary microbial
biofilms were analysed using flow-cytometry and terminal restriction fragment length
polymorphism (T-RFLP) analysis. The structures of the various upcoming communities in the
anode chambers were correlated with pH sensitivity, current production and biofilm formation
exploiting the Dalmatian-Plot and n-MDS similarity analysis.
7.2 Materials and methods
7.2.1 General conditions
All microbiological and electrochemical experiments were conducted under strictly anoxic
conditions at 35°C. If not stated otherwise, all reported pH-values in this study refer to the initial
growth medium pH within the electrochemical cell. All chemicals were of analytical or
biochemical grade. If not stated otherwise, all potentials provided in this article refer to the
Ag/AgCl reference electrode (sat. KCl, 0.195 V vs. SHE). All reported data are based on at least
three independent biological biofilm replicates and two replicates per biofilm for the continuous
flow-mode operation.
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7.2.2 Electrochemical set-up
All electrochemical experiments were carried out under potentiostatic control, using three-
necked-flasks (250 mL) with three electrode arrangement consisting of the working electrode
(projected surface area; 8.00 cm2), a Ag/AgCl reference electrode (sat. KCl, Sensortechnik
Meinsberg, Germany, 0.195 V vs. SHE) and a counter electrode. The working and counter
electrodes used throughout this study were graphite rods (CP-Graphite GmbH, Germany). The
counter electrode was separated from the growth medium by a Nafion® 117 perfluorinated
membrane. The experiments were conducted with a Potentiostat/Galvanostat Model VMP3
(BioLogic Science Instruments, France), equipped with 12 independent potentiostat channels.
Cyclic voltammetry (CV) was performed during turnover and non-turnover conditions at a scan
rate of 1 mV s-1
in accordance with previous studies, e.g. (Fricke, et al., 2008, Srikanth, et al.,
2008). The current density is reported per projected surface area and denominated as “geometric
current density”.
7.2.3 Microbial inoculum and growth medium
The source for the microbial inoculum was primary wastewater collected from the WWTP
Steinhof, Braunschweig (Germany). The pH of the wastewater inoculum was 6.7±0.1 all time.
Always the identical inoculum was used for a consecutive set of experiments (see Section 7.3.3.)
The bacterial growth medium was prepared as reported by Kim et al. (Kim, et al., 2005). It
contained NH4Cl (0.31 g L−1
), KCl (0.13 g L−1
), NaH2PO4•H2O (2.69 g L−1), Na2HPO4 (4.33 g
L−1
), trace metal (12.5 mL) and vitamin (12.5 mL) solutions (Balch, et al., 1979). Acetate (10
mM) served as substrate in the growth medium. The pH values used during this study were 3, 5,
6, 7, 8, 9 and 11. The pH of the growth medium was adjusted to the desired value by using 1 N
NaOH or O-phosphoric acid. In the cathode chamber, buffer solutions were set to an equal pH-
value as the anodic pH and replenished in line with the anode solutions in the fed-batch
experiments. In order to ensure anaerobic conditions the growth medium was purged with
nitrogen at least for 20 min before use.
7.2.4 Biofilm growth (fed-batch experiments)
As described by Liu et al. (Liu, et al., 2008) for the formation of primary biofilms, 6.5 mL of
wastewater were inoculated into the sealed cell, containing 193 mL medium spiked with 10 mM
acetate as carbon and energy source. The cell was operated at 35°C. For consecutive cultivations
(that is exchanging the medium of the anode chamber by removing the old medium refilling it
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with fresh medium and carbon source) 80% of the anode solution was exchanged, the volume
was chosen for practical reasons of reactor operation, but no new inoculation by a wastewater
took place. The exhausted substrate solutions of consecutive replenishing cycles were
denominated as “CX”, e.g. C1 to C5 for a 5 week operation period. During medium
replenishments, the cathode solution was also completely replenished using solutions of
identical pH like in the anode compartment. A constant potential of 0.2 V was applied to the
working electrode to facilitate the formation of a bioelectrocatalytic biofilm. The growth of the
biofilm was monitored by measuring the bioelectrocatalytic oxidation current. The exhausted
substrate was replenished regularly and the substrate level was monitored via HPLC. The data
for the maximum current generation at different constant pH-values (section 7.3.1.) are based on
biofilms grown for at least 4 fed-batch cycles and showing a constant performance (see e.g.
(Liu, et al., 2008)). These biofilms were also used for the flow-through (section 7.3.2.) and
cyclic voltammetric (7.3.3.) experiments. Furthermore selected fed-batch experimental runs
were used for the microbial analysis (7.3.4.).
7.2.5 Biomass determination
At least three independent samples per biofilm (each 1.2 mL) were spinned down at 17900 g for
10 min at 4°C in tubes, which prior to the analyses were dried at 105°C for 24 h. The
supernatant was removed and the procedure repeated until a cell pellet was accumulated (usually
less than 5 times). Subsequently the identical drying procedure was applied and the mass
difference of each tube, representing the dry mass per biofilm sample, was determined.
7.2.6 Metabolic analysis
Acetate consumption was analysed by HPLC (Spectrasystem P400, FINNIGAN Surveyor RI
Plus detector, Fisher Scientific, Germany) equipped with a Rezex HyperREZ XP Carbohydrate
H+ 8 µm column. Chromatograms were recorded at room temperature with 0.005 N sulphuric
acid as eluent.
7.2.7 Continuous flow mode operation and pH-regime studies
Two plastic tanks (10 L each) served as reservoirs for the substrate and buffer solutions. The
flow rates of both solutions were maintained at 0.5 mL min-1
using a peristaltic pump (IP 65,
ISMATEC, Laboratoriumstechnik GmbH, Germany). After adjustment of the biofilms to
continuous flow conditions – represented by establishing a continuous current generation for at
least 12 h - the biofilms were exposed to a pH-ramp from the initial pH-value to pH 5 and then
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to alkaline pH up to pH 11 using a step wise pH decrease/increase with the interval of pH 1 by
changing the influent solution tanks. Thereby the steady-state current at every pH-value was
recorded.
7.2.8 Microbiological analysis
7.2.8.1 Flow-cytometry
Flow cytometry was used to resolve the community structure both in the anode surface and the
replenished substrate solutions on the single cell level. Therefore, every cell in the system was
measured according to the cells’ specific characteristics. These were morphological features
analysed by forward scattering (FSC) and related to cell size and DNA contents which were
specifically stained with the AT specific fluorescent dye DAPI. Every bacterial cell contains at
least one chromosome of a certain length and information. Some cells contain chromosomes of
different length and information whereas most cells contain several copies of them due to the
cells activity state in the cell cycle. After cytometric analysis of these parameters, cells cluster in
distinct patterns within so called dot plots. These patterns represent fingerprint like pattern of a
certain community structure. The patterns are very stable and can easily be reproduced (Müller
and Nebe-Von-Caron, 2010). Changes in community structures can quickly be visualized.
7.2.8.1.1 Sample fixation and DNA staining
Cells were harvested from the wastewater inoculum and from the anodic biofilms formed at pH
6.0, 7.0 and 9.0 and were conserved in fixation buffer (pH 7.0) with 10% sodium azide (Merck,
Germany) dissolved in PBS (1 ml fixation buffer for app. 3 x 108 cells ml-1
) for a maximum of 9
days. Aliquots of the fixed samples were washed twice in 2 ml PBS by centrifugation at 3200 x
g for 5 min and treated gently within an ultrasonic bath for 5 min to dissolve the biofilm. Two
mL of diluted cell suspension were treated with 1 mL solution A (2.1 g citric acid/0.5 g Tween
20 in 100 mL bidistilled water) for 10 min, washed and resuspended in 2 mL solution B (0.68
µM 4‘,6-diamidino-2‘-phenylindole (DAPI, SIGMA), 400 mM Na2HPO4, pH 7.0) and stained
for at least 60 min in the dark at 20°C.
7.2.8.1.2 Multiparametric flow-cytometry
Flow-cytometric measurements were carried out using a MoFlo cell sorter (DakoCytomation,
Fort Collins, CO, USA) equipped with two water-cooled argon-ion lasers (Innova 90C and
Innova 70C from Coherent, Santa Clara, CA, USA). Excitation by 400 mW at 488 nm was used
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to analyse the forward scatter (FSC) and side scatter (SSC) as trigger signal at the first
observation point, using a neutral density filter with an optical density of 2.3. DAPI dye was
excited by 100 mW of ML-UV (333-365 nm) at the second observation point. The orthogonal
signal was first reflected by a beam-splitter and then recorded after reflection by a 555 nm long-
pass dichroic mirror, passage by a 505 nm short-pass dichroic mirror and a BP 488/10. DAPI
fluorescence was passed through a 450/65 band pass filter. Photomultiplier tubes were obtained
from Hamamatsu Photonics (models R928 and R3896; Hamamatsu City, Japan). Fluorescent
beads (Polybead Microspheres: diameter, 0.483 µm; flow check BB/Green compensation Kit,
Blue Alignment Grade, ref. 23520, Polyscience, USA) were used to align the MoFlo (coefficient
of variation – CV value - about 2%). Furthermore, an internal DAPI-stained bacterial cell
standard was introduced for tuning the device up to a CV value not higher than 6%. Cell
aggregation was not observed, thus clearly separated sub-communities were analyzed.
7.2.8.2 T-RFLP and Sequencing
T-RFLP gives information on phylogenetic relationships of bacteria and was therefore used to
prove the presence of certain bacteria on the anode biofilm and within the anode chamber
community. Sequencing was used to certainly affiliate the anode biofilm species to the data
base.
Fixed samples were spinned for 10 min at 17900 g at 4°C and the pellets stored at -20°C until
further analysis. DNA was extracted using a Chelex based method (Giraffa, et al., 2000).
Depending on the pellet size 150 or 300 μL 10% (w/v) Chelex were used. PCR, t-RFLP and
sequencing were performed as described earlier (Harnisch, et al., 2011). The restriction
endonucleases RsaI and MspI (New England Biolabs, Schwalbach, Germany) were used with
the corresponding buffer. Partial sequencing of the 16S rRNA gene was performed with the
primers 27f und 519r and the sequences deposited in the GenBank database under accession
numbers JN393007–JN393010.
The lengths of the fluorescent terminal restriction fragments (T-RFs) in the range from 50 to 600
bp were determined with the genemapper V3.7 software (Applied Biosystems, Weiterstadt,
Germany). Data normalization was performed with an R implementation based on Abdo et al.
(Abdo, et al., 2006) and statistical analysis was done with PAST
(http://folk.uio.no/ohammer/past/).
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7.3 Results and discussion
7.3.1 Biofilm formation and performance at different constant pH
Figure 7-1 summarizes the results of potentiostatic fed-batch experiments of primary,
wastewater derived, biofilms (Liu, et al., 2008, Patil, et al., 2010) for different pH-values during
biofilm formation and operation. It depicts the maximum geometric current densities as well as
coulombic efficiencies of mature primary biofilms grown and operated at different pH-values.
Thereby, Figure 7-1 clearly shows that only at pH 7 a high average current density of 821 µA
cm-2
and coulombic efficiency of 82% was achieved, whereas more acidic or more alkaline
conditions, i.e. pH-values differing from the pH of the municipal wastewater (pH 6.7) that
served as inoculum, resulted in a clear decrease of current density and coulombic efficiency.
Figure 7-1 Performance of electroactive biofilms grown and operated at different pH-values:
Maximum current densities (filled circles; derived from chronoamperometric fed-batch
experiments at 0.2 V vs. Ag/ AgCl) and coulombic efficiencies (open squares) of primary,
wastewater derived biofilms are shown. The substrate was 10 mM acetate.
One can clearly see that at extreme pH-values, i.e. at pH 3 and pH 11, no bioelectrocatalytic
activity was established, whereas the activity at pH neutral is in accordance with previous
studies (e.g., (Patil, et al., 2010)). This result was not unexpected, as the primary wastewater
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from the local treatment plant possessed an almost neutral pH, and thus the microbial
communities therein can be assumed to be properly adapted to this pH-environment.
Interestingly, whereas the average maximum geometric current density, as a measure of the
maximum performance, is only slightly decreasing from pH 7 to 9 (~10%) the coulombic
efficiency, representing the cumulative performance, is more severely affected. It decreases
from 82% at pH 7 via 73% at pH 8 to 39% at pH 9. These results clearly show that – when using
identical inoculums of pH neutral wastewater– electrocatalytic active biofilms can be derived
only in a limited pH-window (here from pH 6 to pH 9).
In conclusion it can be stated that the more the pH-value during biofilm formation and operation
deviates from the pH of the bacterial source (pH neutral wastewater), the lower and less efficient
its bioelectrocatalytic activity becomes.
7.3.2 Biofilm performance at varying pH-environment during operation
Subsequently, the performance response of well developed biofilms on the variation of the pH-
environments was assessed, in order to mimic the influence of a changing pH in the wastewater
influent. This is not only highly relevant concerning the technical applicability of the
electroactive biofilms, but can furthermore be regarded as stress-test from the microbiologist’s
perspective. Figure 7-2A shows the bioelectrocatalytic current production of a mature, i.e.
constant current producing, biofilm (grown at pH 7) in a continuous flow mode reactor (see
7.2.7.) exposed to varying pH-values.
One can clearly see that the bioelectrocatalytic performance declines when exposing the biofilm
to more acidic conditions. For the acidic pH-environment the bioelectrocatalytic performance
drops almost completely down, from about 800 µA cm-2
at pH 7 to less than 40 µA cm-2
at pH
5. Remarkably, as Figure 7-2 A shows, the complete bioelectrocatalytic activity is re-established
within less than 24 h, when switching the pH back to pH 7. Furthermore, Figure 7-2 A shows
that an exposure to more alkaline pH first slightly increases the bioelectrocatalytic current
production. However, longer exposure times and especially highly alkaline conditions lead to an
irreversible biofilm degradation that cannot be re-established when the biofilm is exposed to pH
7 again (see Figure 7-2A).
This biofilm degradation often went along with a biofilm detachment from the electrode surface,
as can already be seen by visual inspection (see Figure S11-11 in Supplementary information).
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Figure 7-2 A) Chronoamperometric current density changes (at 0.2 V vs. Ag/ AgCl) for a
biofilm initially grown at pH 7.0 in relation to variations of the growth medium pH (numbers
indicate the respective pH-value of operation); B) Steady state current densities at 0.2 V vs. Ag/
AgCl of biofilms grown at pH 8 (circles) and pH 7.0 (squares) at varying medium pH (derived
from experiments similar as shown in A)).
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Figure 7-2B summarises the results of similar experiments, as in Figure 7-2A, performed with
biofilms grown at pH 7 and pH 8 , by depicting the respective current density as function of the
applied pH-value. Commonly, the operational window is limited to pH-values between pH 6
and pH 9, which is well in accordance with the pH-window found for the formation of
respective bioelectrocatalytic biofilms from wastewater inoculums (see Figure 7-1).
Furthermore, it can be concluded that pH 7 grown biofilms (showing 360 µA cm-2
) are about
twice as active at pH 6 than biofilms formed at pH 8 (171 µA cm-2
). Thus, one can assume that
these biofilms are better adapted to the respective higher proton concentration. Furthermore,
these results are well in accordance with preceding MFC studies, in which a reversible
adaptation of anodic biofilms to varying pH-conditions, resulting in different reactor
performances, was demonstrated, e.g. (Jadhav and Ghangrekar, 2009).
7.3.3 Influence of the pH and buffer capacity on the electron transfer
In order to elucidate the influence of the anode chamber’s pH and ionic strength/ buffer capacity
on the electron transfer cyclic voltametric measurements (CV) were performed. Consequently,
to minimize the impact of any biological variability in the experiments always identical biofilms
were studied for varying pH-conditions by changing the medium in the anode chamber between
the experiments (overall duration was less than 6 h). Figure 7-3 shows the non-turnover, i.e.
acetate depleted, CVs of a pH 7 grown biofilms at different pH-values in pure electrolyte
solutions.
The correlation between pH-value and redox-potentials of the active sites depicts that with
decreasing pH the formal potentials of all redox-active moieties are shifted towards more
positive values. The redox-centres, most likely related to the direct electron transfer sites of
Geobacter (Fricke, et al., 2008, Liu, et al., 2008) as these are the dominating microorganism in
this biofilm (see below) are both ascribed to c-type cytochromes (Millo, et al., 2011), shifting
more than 140 mV from pH 9 to pH 6. This potential shift of about 47 mV/ pH is well in
accordance with previous studies for G. sulfurreducens (that was a dominant microorganism in
our biofilms, see 7.3.5.) on glassy carbon electrodes (Katuri, et al., 2010) and acetate derived
biofilms (Yuan, et al., 2011), both studied for pH 6 to pH 8. Interestingly, when biofilms were
exposed to pH 5, which after a longer exposure was generally leading to a complete biofilm
detachment, no constant and distinctive CV-curve could be recorded.
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Figure 7-3 Influence of the operational pH: Cyclic voltammograms obtained at different
operation pH (using a constant ionic strength of 50 mM) at a scan rate of 1 mV s-1
during non-
turnover conditions for wastewater derived, acetate-fed biofilm formed at pH 7.0. (For pH 6 to
pH 8 steady-state CVs are shown, for pH 5 the 3rd CV-curve).
Analysing both oxidation peaks of the direct electron transfer proteins showed a lower
discrimination between the redox-couples at more acidic conditions (data not shown). This
finding, indicating a different pH-dependence of both direct electron transfer sites, needs a more
detailed analysis applying highly-sensitive electroanalytical methods as well as hyphenated
techniques, e.g. spectroelectrochemical approaches, e.g. (Millo, et al., 2011).
Interestingly, the more positive oxidation peak, located at about -100 mV at pH 7, which is not
involved in the bioelectrocatalysis (Fricke, et al., 2008), shows also a pH dependence.
Subsequently, in order to elucidate if the demonstrated dependence of the electron transfer is
purely associated to H+-transfer or depends on a charge balancing counter ion transfer in
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general, CVs for constant pH but varying buffer capacity were recorded (Figure S11-1A). These
show clearly that a variation of the buffer capacity has almost no influence on the formal
potentials of the active site (Figure S11-1A). Only a decreasing CV-resolution, which might be
attributed to the ohmic resistance (i×R-drop) in the biofilm, can be detected. Thus, the buffer
capacity (and ionic strength) has no influence on the electron transfer thermodynamics.
In contrast, the buffer capacity determines the maximum current density for turnover conditions,
i.e. the maximum bioelectrocatalytic performance. An increase by one order of magnitude in
buffer concentration caused an increase of the maximum current production of about 40% for
the identical biofilm (see Figure S11-1B: 250 µA cm-2
and 400 µA cm-2
were achieved for 5
mM and 50 mM buffer concentration, respectively).
This finding is well in accordance with previous studies showing the pH gradient within a
biofilm from the electrode surface to the electrolyte solution is severely limiting the
bioelectrocatalytic performance (Torres, et al., 2008).
These results clearly reveal that the charge balancing ion (proton) transfer through the biofilm
represents a severe bottleneck of the electrocatalytic biofilm activity. This was already indicated
in prior CV-studies showing a differing mass-transfer dependence of these biofilms for low and
high scan rates (Fricke, et al., 2008, Srikanth, et al., 2008) and recent results mapping the pH-
gradient within G. sulfurreducens biofilms (Franks, et al., 2009).
7.3.4 Microbial biofilm analysis
In order to elucidate the microbiological reasons for the variations in the bioelectrocatalytic
performances of the natural community derived biofilms gained at different pH-values, flow-
cytometric and T-RFLP-analyses including 16S rRNA gene sequencing were performed. In
total, two out of several parallels, in the following denominated as electrode-set 1 and electrode-
set 2, were investigated on their microbial structure (by flow-cytometry) and composition (by T-
RFLP) at pH 6, pH 7 and pH 9 using the identical inocula of pH 6.7 wastewater for the
respective parallels and acetate as carbon and energy source.
Figure 7-4 shows, on the example of a pH 7 derived biofilm at electrode-set 1, the flow-
cytometric analysis of a bacterial anode chamber community after wastewater inoculation. Five
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successive fed-batch medium exchanges were performed (denominated as C1 to C5, see
materials and methods) and the dynamics of the planktonic anode chamber community followed
until final development of the active current producing anodic biofilm. The resulting datasets
showed the clustering of cells of the community to distinct sub-communities (Müller and Nebe-
Von-Caron, 2010). Presence and absence as well as the relative abundances of cell numbers
within these clusters gave a fingerprint like information on the structure of the microbial
community. It is obvious that the structure was changing over the five feeding cycles C1 to C5
which is due to the adaptation of the community structure from complex wastewater to acetate
as sole carbon and energy source. It shows that the microbial community responded sensitively
to changes in its microenvironments (Günther, et al., 2011).
Additionally, the cytometrically determined structures of the communities in the anode chamber
during the different feeding cycles differed strongly from that of the inoculum (see Figure 7-4).
Despite the miscellaneous structure variation in the chamber broth (flow-patterns C1 to C5), a
microbial biofilm evolved at the anode dominated by mainly one phylotype.
For both electrode-sets a similar maximum current density and columbic efficiency for biofilms
formed at pH 7 was achieved, with in average jmax=740 µA cm-2
and CE=99.8%. The anode
biofilms of both electrode-sets were dominated by one phylotype, as can be seen from the
respective flow-cytometric analysis. The related T-RFLP chromatograms for restriction
digestion analysis displayed only one dominant peak at 238 bp (92% of the total peak area for
pH 7) (see S11-2). The subsequent 16S rRNA PCR products of the two investigated pH 7 anode
biofilms were partially sequenced and resulted in the identification of the genus Geobacter using
the RDP classifier. The maximum score in the BLAST search, excluding
uncultured/environmental samples, resulted in the identification of Geobacter sulfurreducens
(CP002031.1) with a maximum identity of 97%. Comparison of these sequences with previous
results in the work group shows a 100% identity (Harnisch, et al., 2011). This dominance of
Geobacter sulfurreducens for the respective conditions is well in line with a previous study
(Torres, et al., 2010).
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Figure 7-4 Bacterial community profiles of the inoculum and the successive media of the anode
chamber of a pH 7 grown biofilm (electrode-set 2). The profile of the community is
cytometrically determined by the cells’ DNA content labelled with the A-T specific fluorescent
dye DAPI and the cells’ forward scatter behaviour (FSC). As a result fingerprint-like cytometric
patterns emerged as subsets of cells which gather in numerous clusters of changing cell
abundances therein. Up to 250000 cells were analysed and the dominant sub-populations
presented in yellow colour. The peak in the lower left corner of the histograms represents the
noise of the cytometer as well as unstained cell debris.
As was shown for both of the electrode-sets and for the three different pH-values investigated
the community structure responded with rapid and dynamic changes in community structure
since it adapted easily to the varying abiotic conditions. To align fingerprints originating from
‘healthy’ and active biofilm launching communities apart from those connected with inactive
biofilms a similarity analysis was performed on the basis of the cytometric dot plot clusters. The
approach divides the productive from the non-active communities in the sense of their potential
to establish active biofilms with high current production efficiency. Thus, cytometric analysis
gives information on the potential ability of a community to set up active biofilms. The
approach is quickly performed and cheap. This is a huge advantage in comparison to
phylogenetic approaches like T-RFLP and sequencing techniques which are considerably more
cost and time intensive.
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The n-MDS (see Section 11.9 in Appendix C) analysis of the anode chamber microbiology
revealed that the planktonic communities and the biofilms are changing gradually in their
structure and dynamics depending on experimental conditions. The pH-environment seems to
influence the planktonic community structure more distinct than the consecutive acetate fed-
cycle conditions since all of the latter are more or less closely clustered (high similarity). This
finding points towards a high stability of the planktonic microbial community over longer time
periods (more than 2 weeks) under stable pH and substrate (acetate) conditions. However,
changes in the pH caused also changes in the planktonic community structure, clearly shown by
the separated respective Dalmatian plots. The related consecutive fed-cycles at the various pH-
values did not cluster together (low similarity). Evaluating the anode biofilm patterns at pH 6,
pH 7 and pH 9 pronounced differences in the Dalmatian pattern were identified, resulting in a
clear separation from all planktonic community plots (low similarity).
Furthermore, high performing biofilms, identified to contain mainly Geobacter sulfurreducens
clustered together in the lower right corner of the n-MDS plot (electrode-sets 1 and 2, pH 7;
electrode-set 2, pH 6; electrode-set 1, pH 9, high similarity). The patterns of the low performing
electrodes (electrode-set 2, pH 9; electrode-set 1, pH 6) showed a different fingerprint and
clustered apart from the other electrodes but also from the planktonic communities (low
similarity).
The opposite development of high performing biofilms at pH 6 and pH 9 can be explained by
the different inocula used for the two electrode-sets 1 and 2. The different inocula were obtained
during either a summer (August) and winter (November) period from the wastewater treatment
plant and influenced the biofilm establishment insofar that either pH 6 (electrode-set 1) or pH 9
(electrode-set 2) resulted in a microbial underdevelopment represented by an only low
bioelectrocatalytic activity.
The two pH 7 biofilms showed a constant high bioelectrocatalytic activity and typical
population patterns for G. sulfurreducens, as measured by flow-cytometry. The further two high
performing biofilms found for pH 6 and pH 9 (with performance values of jmax=705 µA cm-2
and CE=50.2% for pH 9 (electrode-set 1) and jmax =191 µA cm-2
and CE=72.1% for pH 6
(electrode-set 2)) showed similar population patterns to the pH 7 grown biofilms (see Figure 7-
5). Here, the T-RFLP data presented the respective peak at 238 bp (84% of the total peak area
for pH 6 grown biofilms, Fig S11-4 (electrode-set 2); for pH 9 grown biofilms, S11-3
(electrode-set 1)) which also confirmed G. sulfurreducens using partially sequencing.
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In contrast, when the biofilms possessed only a lower electrochemical activity degenerated
cytometric biofilm patterns were observed (more distant clustering in Figure 7-5); jmax=137 µA
cm-2
and CE=21% for pH 6 (electrode set 1) and jmax=0.27 µA cm-2
and CE=0.1% for pH 9
(electrode set 2). For the latter, also a decrease in anode dry biomass from 29.8 mg ± 0.2 mg (pH
7) to 9.8 mg ± 0.2 mg (pH 9) was detected. In both cases of low-performance biofilms G.
sufurreducens was not the dominating organism of the respective anode community (see S11-3
(pH 9, electrode set 2) and S11-4 (pH 6, electrode set 1)).
Thus, since substrate and electrode potential were identical for all experiments it is obvious that
the pH-environment as well as the microbial source were the strongest driving forces for
variations within the bioelectrocatalytic activity on the anode and the structures of the associated
planktonic microbial communities. These findings suggest that current production and efficiency
depend on a stable anode biofilm formation, allowing (for the applied conditions in this study)
the dominant growth of G. sufurreducens. The varying biofilm formation efficiency at different
pH-values is not surprising since G. sulfurreducens is known to possess its maximum growth
rate at pH 7 that is significantly lowered, e.g. at pH 6 (Franks, et al., 2009) - which is in
accordance with the DNA/FSC growth pattern detected via flow-cytometry.
Wastewater is highly variable in its composition and characterized by inconstant abiotic
parameters. How sensitive communities react to extrinsic parameters like wastewater
composition and temperature is shown within the Dalmatian plot with the two inocula clustering
apart from each other in their community pattern but also apart from the newly developed
communities in the anode chamber now using only acetate as substrate. Therefore it can be
assumed that wastewater composition and thus its microbial community severely influences the
ability to form a high performance biofilm at the respective pH-value other than the growth
optimum, as the environmental conditions and their variability (frequency and amplitude) define
which microbial species can establish and persist (Günther, et al., 2011). Furthermore, it has to
be pointed out that no other single microbial species dominated the low performance biofilms,
which in contrast where highly diverse. Here, different, complex ecological mechanisms,
including ‘priority effects’ and the formation of repellent EPS substances, may avoid the
dominance of a distinct phylotype for the respective operating conditions (Flemming and
Wingender, 2010, Van Gremberghe, et al., 2009).
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Although T-RFLP pattern suggested a complex suspension community in the anode
compartment of all biofilms, there was seemingly no other bioelectrocatalytic microorganism
present that could take advantage and dominate the biofilm instead of Geobacter sulfurreducens.
Figure 7-5 Dalmatian-n-MDS analysis with overlaid cytometric flow-plots derived from anode
chamber communities and anode biofilms when treated over several feeding cycles and different
pH-values. Black patches in flow-plots depict gate positions, cycle number is given with c 1–5
and pH-affiliation with various grey/black labels (black: pH 7, grey: pH 9, light grey: pH 6, bold
fringe around flow-plot: electrodes; details see text and S11-2 to S11-10 for raw data).
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7.4 Conclusions
It is demonstrated that the pH-value plays an important role for the biofilm formation,
composition and performance of electroactive biofilms. Starting from pH-neutral wastewater as
inoculum, high performance biofilms were dominated by G. sulfurreducens, whereas a lower
performance went along with a higher microbial diversity. Analysing the impact of the pH-value
and buffer capacity/ ionic strength on identical biofilms shows that the latter influence the
maximum achievable current density, but not the formal potentials of the electron transfer
proteins. These, however, show a strong dependence on the pH-value of the solution - calling
for the further investigation of this phenomenon.
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CHAPTER VIII
8 Electroactive mixed culture derived biofilms in microbial
bioelectrochemical systems: the role of inoculum and
substrate on biofilm formation and performance
8.1 Introduction
Bioelectrochemical systems (BESs) are a group of developing and promising technologies
targeting different kind of goals (Rabaey and Rozendal, 2010), from the production of
bioelectricity, via the production of biofuels (e.g., H2), to the production of valuable
biochemicals (e.g., H2O2). As seen in chapter 1, depending on the BES’s application (see Fig. 1-
6), a plenitude of applications can be conceived regarding the overall configuration of the BES
(Logan, et al., 2008, Logan, et al., 2006), from the membrane specificity (Harnisch and
Schröder, 2009) to the type of (bio) catalyst interacting at both electrodes (Franks, et al., 2010,
Rosenbaum, et al., 2011). BESs utilize the energy available in bio-convertible substrates via the
catalytic activity of electrochemically active biofilms developed at the electrode material (in this
case the anode). These biofilms are composed of a network of bacteria layers growing on the
electrode material that oxidize a substrate to finally transfer the harvested electrons to the anode
that serves as a microbial electron acceptor (Lovley, 2011).
Commonly in BESs the focus in using pure cultures of bacteria as bio-catalyst is the study of
fundamental phenomena such as the thermodynamic processes involved in microbial electron
transfer (see Chapter 2, 3 and 4). However most BESs take advantage of mixed culture derived
anodic biofilms due to their practical application. To find the optimal conditions for the
formation and performance of electroactive biofilms, different microbial sources and type of
substrates should be explored to get a more precise idea of the dynamics of the electrode
bacterial communities (Logan and Regan, 2006a, Pant, et al., 2010). It has been demonstrated
that many different factors influence both, the composition of the bacterial community in
electroactive biofilms in BESs and their performance (Franks, et al., 2010). Some of the studied
factors are, among others: temperature (Patil, et al., 2010), external ohmic resistance (Zhang, et
al., 2011), oxygen limitation (Biffinger, et al., 2009), flow rate (Ieropoulos, et al., 2010), pH
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environment (see Chapter 7), type of inoculum and type of substrate as it has been explored in
this study.
Regarding the influence of the inoculum and substrate on the formation and performance of
anodic biofilms, one can find in the literature the following representative examples exploring
the influence of certain inocula and substrates. Min, et al. (Min, et al., 2005) compared two
types of inocula for power production: a pure culture of Geobacter metallireducens and a mixed
culture enriched from wastewater that commonly leads to the formation of a biofilm dominated
by a bioelectroactive bacteria, strain of the Geobacteraceae family (Harnisch, et al., 2011).
Since both biofilms were possibly conformed by similar electroactive bacteria is not surprising
that no significant difference was found in the performance between used inocula.
A similar approach was used by Ieropoulos, et al. (Ieropoulos, et al., 2010). In their research
they tested two inocula representative of complex communities of microflora found in the final
stages of wastewater treatment (anaerobic and aerobic effluent), an environmental inoculum
such as river water and a pure culture of a bioelectroactive bacteria such as G. sulfurreducens.
The study by Ieropoulos et al. showed no significant difference in the performance no matter
what inoculum was used. Furthermore although they report the characterization of the anodic
biofilm, no quatitative data were presented to allow a proper comparison. On the other hand,
Nimje et al. (Nimje, et al., 2012) tested two different inocula (wastewater and a pure culture of
Shewanella oneidensis MR-1) and four types of wastewaters as substrate sources (agriculture,
domestic, paper and food/dairy). Their study produced results which corroborated the findings
of a great deal of the previous BES experiments using different kind of inocula and substrates,
i.e., there was no significance difference in the performance of the tested systems. This probably
due to the mixture of a pure culture such as S. oneidensis MR-1 with wastewater as inoculum,
which probably led to the dominance of electroactive bacteria present in the wastewater and thus
masking the effect of S. oneidensis MR-1.
Additionally, there are a few studies focused on the influence of different substrates on the
biofilm formation. Velasquez-Orta et al. (Velasquez-Orta, et al., 2009) tested two kind of algae
as substrate for BESs. They demonstrated that in principle algae can be used as a renewable
source of electricity production. However they could not find significance differences by
feeding different kind of algae. From the different substrates used in comparative BES studies,
carbohydrates are the most used due to the preference of some electroactive bacteria for these
compounds (e.g., acetate in the case of Geobacter and lactate in the case of Shewanella). Several
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research groups have studied the influence of some carbohydrates. For example Min and Logan
(Min and Logan, 2004) studied the influence of five specific substrates (glucose, acetate,
butyrate, dextran and starch) finding that when the system was fed with acetate the power
generation was sustained at high rates. In a similar study performed by Thygesen et al.
(Thygesen, et al., 2009) several BESs were fed with acetate, glucose or xylose as substrates.
Acetate produced the highest current probably due to a simpler metabolism than with glucose or
xylose. In another major study, Lee et al. (Lee, et al., 2008) quantified the impact of using
acetate and glucose as substrates on several experimental variables such as current and biomass
production, among others. The energy-conversion efficiency was significant higher with acetate
than with glucose. They attributed this to very low energy-conversion efficiency for glucose due
to a large increase of the anode potential. Additional analysis of the biomass on the anode
showed that although glucose allowed higher biomass density, it had a very low current density,
which supported the fact that the density of electroactive bacteria was very low.
Although acetate seems to be the best substrate for electroactive biofilms enriched from
wastewater samples, some recent studies show the contrary. For instance Cao et al. (Cao, et al.,
2010) demonstrated that by using three specific substrates like glucose, acetate and ethanol for
the growth of electroactive biofilms glucose is utilized in a more efficient way to produce
current than the rest of substrates. Results presented by Cao et al. (Cao, et al., 2010) were in
agreement with similar studies published by Sharma and Li (Sharma and Li, 2010) showing the
same trend in the utilization of different substrates by electroactive bacteria.
In a different category, experiments using a co-culture of bacteria which benefit from their
interaction should be mentioned because this type of studies allow us to understand the
ecological relationships of the microbiota in BESs, a necessary requisite to gain deeper insight
into their performance. For instance, Venkataraman and co-workers (Venkataraman, et al.,
2011) showed that the fermentation product 2,3-butanediol stimulates mutually beneficial
interactions between Pseudomonas aeruginosa PA14 and Enterobacter aerogenes in a BES
with glucose as the initial substrate under microaerobic conditions. They found that current
density by a co-culture of P. aeruginosa and E. aerogenes increased at least 14-fold compared to
the current density by either of these two bacteria alone; and that E. aerogenes fermented
glucose principally to 2,3-butanediol, which was subsequently consumed by P. aeruginosa. The
current production by a pure culture of P. aeruginosa with 2,3-butanediol was increased 2-fold
compared with glucose as the carbon source. This was due to enhanced phenazine production by
P. aeruginosa. Their study was the first to demonstrate metabolite based ‘‘inter-species
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communication’’ in BESs, resulting in enhanced electrochemical activity. It also explains how
an inconsequential fermenter can become an important electrode respiring bacterium within an
ecological network at the anode.
As one can see, it exists a vast amount of BES studies using all kind of inocula and substrates,
however there is a lack of unifying studies that compare different inocula and substrates under
the same experimental conditions. Thus, in order to exclude the influence of operational
variables and to investigate only the effect of individual microbial inoculum source with sodium
acetate or sodium lactate as substrates, the experiments here presented were conducted with
half-cell set-ups under potentiostatic control (Fig. 8-1) investigating the general
bioelectrocatalytic activity (current density) and the voltammetric behavior.
8.2 Materials and methods
8.2.1 General conditions
All microbiological and electrochemical experiments were conducted under strictly anoxic
conditions at 35°C. All chemicals were of analytical or biochemical grade. If not stated
otherwise, all potentials provided in this article refer to the Ag/AgCl reference electrode (sat.
KCl, 0.195 V vs. SHE). All data are based on experiments during at least 5 semi-batch cycles of
2 independent biofilm replicates, and the standard deviations are presented in Fig. 8-2.
8.2.2 Electrochemical set-up
All electrochemical experiments were carried out under potentiostatic control, using three-
necked-flasks (250 mL) with three electrode arrangement consisting of the working electrode
(projected surface area; 8.00 cm2), a Ag/AgCl reference electrode (sat. KCl, Sensortechnik
Meinsberg, Germany, 0.195 V vs. SHE) and a counter electrode. The working and counter
electrodes used throughout this study were graphite rods (CP-Graphite GmbH, Germany)
contained in the same chamber (Fig. 8-1A). The experiments were conducted with a
Potentiostat/Galvanostat Model VMP3 (BioLogic Science Instruments, France), equipped with
12 independent potentiostat channels. The current density is reported per projected surface area
and denominated as “geometric current density”.
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Figure 8-1 A) Electrochemical half cell set-up under potentiostatic control and B) Exemplary
established bioelectrochemical active biofilm enriched from primary wastewater fed with
acetate. The red color is mainly caused by the hemes (Jensen, et al., 2010).
8.2.3 Microbial inoculum and growth medium
There were four types of wastewater collected from the Waste Water Treatment Plant Steinhof,
Braunschweig (Germany) that served as the source for the microbial inoculum, i.e.: primary
wastewater, activated sludge, primary sludge and secondary sludge. Always the identical
inoculum was used for a consecutive set of experiments. The bacterial growth medium was
prepared as reported by Kim et al. (Kim, et al., 2005). It contained NH4Cl (0.31 g L−1
), KCl
(0.13 g L−1
), NaH2PO4•H2O (2.69 g L−1
), Na2HPO4 (4.33 g L−1
), trace metal (12.5 mL) and
vitamin (12.5 mL) solutions (Balch, et al., 1979). Sodium acetate (10 mM) or Sodium lactate
(10 mM) served as substrates in the growth medium. In order to ensure anaerobic conditions the
growth medium was purged with nitrogen for 30 min before use.
8.2.4 Biofilm growth in bioelectrochemical half-cells
The biofilm formation procedure was followed as described by Liu et al (Liu, et al., 2008) in
fed-batch experiments. For the biofilm formation 10 mL of individual microbial sample was
inoculated into the sealed electrochemical cell that contained 200 mL of the stirred growth
medium with substrate under study. A constant potential of +0.2 V was applied to the working
electrode to facilitate the biofilm formation. The biofilm growth was monitored by measuring
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the bioelectrocatalytic oxidation current. For the initial (usually three) batch cycles, microbial
inoculum was added to the medium.
8.2.5 Cyclic voltammetry
Cyclic voltammetry (CV) was performed during turnover and non-turnover conditions in
accordance with previous studies, e.g. (Fricke, et al., 2008, Srikanth, et al., 2008). Potentials
were applied from -500 to +300 mV (vs. Ag/AgCl) at a scan rate of 1 mV s-1
with continuous
monitoring of the current response.
8.2.6 Metabolic analysis for coulombic efficiency calculation
Acetate and lactate consumption was analysed by HPLC (Spectrasystem P400, FINNIGAN
Surveyor RI Plus detector, Fisher Scientific, Germany) equipped with a Rezex HyperREZ XP
Carbohydrate H+ 8 µm column. Chromatograms were recorded at room temperature with 0.005
N sulphuric acid as eluent. The total coulombic efficiency (CE) was calculated by integrating
the current over time according to the method described by Logan et al. (Logan, et al., 2006).
8.3 Results and discussion
8.3.1 Current density production of enriched microbial electroactive biofilms as a
function of microbial inoculum and substrate
A significant difference in current generation was observed for all bioelectrochemical set-ups
(Fig. 8-2). However only visible biofilms were detected after 5 semi-batch cycles for
experiments using primary wastewater as inoculum (see Fig. 8-1B). As shown in Fig. 8-2, the
acetate-fed-reactor with primary waste water inoculum showed the highest current density (558
± 27 μA cm-2
, CE = 94 ± 1%), followed by lactate-fed-reactor with primary waste water
inoculum (460 ± 54 μA cm-2
, CE = 25 ± 12%). Secondary sludge resulted in the lowest current
outputs with both substrates in comparison to other inocula. The most possible reason for the
low current densities with secondary sludge as an inoculum could be the absence of high-
current-producing exoelectrogenic microorganisms to develop biofilms either through
competition with other microbes or an inability to use this specific substrate (Lee, et al., 2008).
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Figure 8-2 Bioelectrocatalytic performance of electroactive microbial biofilms derived from
different inocula with fed batch operation in potentiostatically controlled half-cell experiments
(+0.2 V vs. Ag/ AgCl) at graphite rod electrodes. The substrate was 10 mM sodium acetate or
sodium lactate respectively.
Activated sludge as inoculum although did not show as good performance as primary
wastewater, it was the second best current density producer amongst other microbial inocula
with 94 ± 3 μA cm-2
for acetate and for 165 ± 31 μA cm-2
lactate. Furthermore, the high
performance with primary wastewater for the formation of bioelectroactive biofilms
demonstrated its ability as efficient microbial inoculum source. The better performance with
primary waste water also indicated selective enrichment of electrocatalytically active microbes
on the anode and thus proved to be better candidates for the formation of mixed culture based
electroactive bacteria (Harnisch, et al., 2011).
Just two of the used microbial inocula showed significant current density production (primary
wastewater and activated sludge). This could be attributed to the complexity of these mixed
culture inocula (Angenent, et al., 2004, Logan and Regan, 2006a). As shown here and in
accordance with previous results (Chae, et al., 2009, Jung and Regan, 2007, Liu, et al., 2004,
Pant, et al., 2010), acetate was the preferred substrate for electricity generation with different
inocula in MFCs.
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The low performance with primary and secondary sludge might be attributed to a low extent of
bacterial adhesion to the anode which is necessary for better performance. It has been shown
that electroactive biofilm formation at the anodes is an important factor to increase the current
production (Jiang, et al.). Furthermore, these microbial inocula contain a variety of non-
electrogenic bacteria that compete with electrogenic bacteria for the growth, which probably
slowed down the electroactive biofilm formation process and thus the overall bioelectrocatalytic
performance (Wagner, et al., 2002). Interestingly, Activated sludge exhibited better performance
with lactate than with acetate which might be because of involvement of lactate utilizing
microorganisms in this inoculum (Liu, et al., 2007).
8.3.2 Bioelectrocatalytic activity of enriched microbial electroactive biofilms as a
function of microbial inoculum and substrate
Cyclic voltammetry was performed for all established bioelectroactive biofilms formed from
four different inocula and fed either with sodium acetate (Fig. 8-3) or with sodium lactate (Fig.
8-4). Cyclic voltammograms during non-turnover (A, C, E and G in Fig. 8-3 and 8-4) and
turnover (B, D, F and H in Fig. 8-3 and 8-4) conditions with all set-ups confirmed the biofilm
associated current generation. Exemplary CVs of primary wastewater, activated sludge, primary
sludge and secondary sludge based electroactive biofilms indicated the different electro-
chemical behaviour with both substrates. As pointed out in section 8.3.1, only visible mature
biofilms were detected with primary wastewater as inoculum (see Fig. 8-1B). Maturity of
biofilms was confirmed from a constant maximum of current density production and a non-
changing CV shape after the third semi-batch cycle. For turnover CVs of biofilms enriched from
primary wastewater the formal potential of the active site (bioelectrocatalysis) was about -282
mV vs. Ag/ AgCl for sodium lactate fed biofilms and -248 mV for sodium acetate fed biofilms
(derived from the first derivative of CVs for turnover conditions in Fig. 8.5). This clearly
indicates that the used inocula considerably influenced the enrichment of electrochemically
active bacteria with different substrates.
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Figure 8-3 Exemplary cyclic voltammograms. Electroactive microbial biofilms derived from
different inocula grown with Sodium acetate (10 mM) recorded during non-turnover (A, C, E
and G) and turnover conditions (B, D, F and H) conditions. The scan rate used was 1 mV s-1
.
Furthermore, the CV patterns during non-turnover conditions with all inocula (A, C, E and G in
Fig. 8-3 and 8-4) showed very different and complex redox behaviour as well and thus electron
transfer thermodynamics. After the third semi-batch cycle only biofilms enriched from primary
wastewater showed a non-changing CV shape (Fig. 8-3 and 4A). For these non-turnover CVs
the formal potential of the active site (bioelectrocatalysis) was about -300 mV vs. Ag/ AgCl in
agreement with previous results with electrodes modified with G. sulfurreducens biofilms
(Fricke, et al., 2008). This clearly indicates that the used inoculum considerably influenced the
enrichment of electrochemically active bacteria with different substrates. Furthermore, this also
demonstrates that the used conditions lead to the enrichment of a well known electroactive
bacteria (G. sulfurreducens) supporting the observed CV shapes for both, turnover and non-
turnover CVs (Harnisch, et al., 2011).
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Figure 8-4 Exemplary cyclic voltammograms. Electroactive microbial biofilms derived from
different inocula grown with Sodium lactate (10 mM) recorded during non-turnover (A, C, E
and G) and turnover conditions (B, D, F and H) conditions. The scan rate used was 1 mV s-1
.
The fact that no clear CV shape was found for the rest of the inocula could indicate that in those
electrodes there was no a predominant bacteria in the biofilm but an association of different
microbes part of a mixed culture community.
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Figure 8-5 Exemplary cyclic voltammograms from electroactive microbial biofilms derived
from primary wastewater grown with 10 mM sodium lactate (A) or 10 mM sodium acetate (B)
recorded during turnover conditions. First derivatives of biofilms grown with sodium lactate (C)
or sodium acetate (D).
8.4 Conclusions
Within this study it is demonstrated the importance of the inoculum and the substrate selection
by analyzing the current production and the formal potentials extracted from cyclic
voltammetry. By this selection, using acetate and lactate-based artificial wastewater as the
bacterial growth medium and real wastewater as inoculum, cyclic voltammograms shapes
similar to pure culture biofilms of G. sulfurreducens were gained (Fricke, et al., 2008). This
raises the question on how distinctive environmental variables (e.g. the bacterial source and the
substrate) influence the bacterial biofilm composition and dynamics. These and further follow-
up questions are under investigation with flow-cytometry, allowing a high-throughput
characterization of natural microbial communities without any previous knowledge on the
bacterial composition (Harnisch, et al., 2011). The monitoring of microbial communities will
use flow cytometric analyses of cellular DNA and polyphosphate to create patterns mirroring
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dynamics in community structure after the study performed by Günther and co-workers
(Günther, et al., 2011). Additionally, the study will use biostatistics to determine the kind and
strength of the correlation between the presence and abundances of initial and developed
microbial communities. Finally, the bacterial composition of certain subcommunities will be
determined by cell sorting and phylogenetic tools like T-RFLP. Due to the above, the
application of flow-cytometry to electrocatalytic biofilms paves the way to follow the
community dynamics as well as bacterial activity states in response to micro-environmental
changes in high through-put BESs.
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9 Supplementary information: Chapter II
Table S9-1 Comparison of geometric current densities for Shewanella oneidensis Wild-type in
different studies.
jmax-CA/ µA cm-2
Applied E/ V* Ref.
7.9 +0.2 This study
5.1 +0.4 (Babauta, et al., 2011)
2.9 0 (Babauta, et al., 2011)
2.0 +0.2 (Okamoto, et al., 2011)
45.0 +0.2 (Rosenbaum, et al., 2010a)
18.5 +0.043 (Coursolle, et al., 2010)
10.0 -0.195 (Peng, et al., 2010b)
9.7 -0.195 (Peng, et al., 2010a)
17.8 +0.041 (Baron, et al., 2009)
16.0 +0.041 (Marsili, et al., 2008a)
22.9 +0.5 (Cho and Ellington, 2007)
23.6 +0.35 (Cho and Ellington, 2007)
24.3 +0.2 (Cho and Ellington, 2007)
25.7 0 (Cho and Ellington, 2007) *Average maximum current density
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Figure S9-1 Schematic drawing of an electrochemical cell for the study of the electron transfer
mechanisms and current production. The electrochemical cell consists of an anode, a cathode
and, a membrane separating both. An oxidation process occurs at the anode, in this case lactate
oxidation, in which electrons and protons are produced. The electrons flow to the cathode
through an external circuit or potentiostat in which the electrons can be can be quantified.
Meanwhile the protons are released to the media and lately they migrate to the cathode chamber
to react with molecules of water and electrons finally producing hydrogen for example. Figure
drawn with modifications after (Rabaey and Verstraete, 2005, Schröder, 2008).
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Figure S9-2 Electrochemical half cell set-up under potentiostatic control. Description: “Top
view” shows the 5 necks of the 250 mL flask. In section A-A’ details of the working electrode,
counter shielded electrode and reference electrode are given. In section B-B’ the port for
filtrated air, filtrated nitrogen and media supply are detailed.
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Figure S9-3 Exemplary fed-batch chronoamperometric cycles (0.2 V vs Ag/AgCl) of
Shewanella oneidensis MR-1 Wild-type and knock-out mutants on equally-sized graphite rod
anode electrodes, in half cells utilizing lactate (18 mM) as the electron donor and anodes as
electron acceptors.
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Figure S9-4 Cyclic voltammetry at 1 mV s-1
(A, C and E) and First derivative plots of CV data
(B, D and F) of S. oneidensis Wild-type (E and F) and mutants (A and B: ΔpilM-Q/ΔmshH-Q; C
and D: ΔpilM-Q) during Turnover conditions. OxT states for oxidation turnover peak, RedT
states for reduction turnover peak and ET states for redox turnover system. Every time 4
exemplary CVs are shown.
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Figure S9-5 Continuation of Fig. S9-4. Cyclic voltammetry at 1 mV s-1
(G, I and K) and First
derivative plots of CV data (H, J and L) of S. oneidensis mutants (G and H: ΔmshH-Q; I and J:
Δflg; K and L: ΔmtrC/ΔomcA) during Turnover conditions. OxT states for oxidation turnover
peak, RedT states for reduction turnover peak and ET states for redox turnover system. Every
time 4 exemplary CVs are shown.
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Figure S9-6 Cyclic voltammetry at 1 mV s-1
(A, C and E) and First derivative plots of CV data
(B, D and F) of S. oneidensis Wild-type (E and F) and mutants (A and B: ΔpilM-Q/ΔmshH-Q; C
and D: ΔpilM-Q) during Non-turnover conditions. OxT states for oxidation turnover peak, RedT
states for reduction turnover peak and ET states for redox turnover system. Every time 4
exemplary CVs are shown.
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Figure S9-7 Continuation of Fig. S9-6. Cyclic voltammetry at 1 mV s-1
(G, I and K) and First
derivative plots of CV data (H, J and L) of S. oneidensis mutants (G and H: ΔmshH-Q; I and J:
Δflg; K and L: ΔmtrC/ΔomcA) during Non-turnover conditions. OxT states for oxidation
turnover peak, RedT states for reduction turnover peak and ET states for redox turnover system.
Every time 4 exemplary CVs are shown.
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Figure S9-8 Data analysis for each catalytic centre (redox-system I and II). On the left column
an exemplary turnover CV for each strain can be seen. In the center is its respective non-
turnover CV. On the right column the final subtracted CV is presented on which the signal
height of each catalytic wave was estimated at suitable fixed potentials. A-C) ΔpilM-Q/ΔmshH-
Q. D-F) ΔpilM-Q. G-I) Wild-type. (see also Fig. 2-5 in Chapter II for details)
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Figure S9-9 Continuation of Fig. S9-8. Data analysis for each catalytic centre (redox-system I
and II). On the left column an exemplary turnover CV for each strain can be seen. In the center
is its respective non-turnover CV. On the right column the final subtracted CV is presented on
which the signal height of each catalytic wave was estimated at suitable fixed potentials. J-L)
ΔmshH-Q. M-N) Δflg, P-R) ΔmtrC/ΔomcA. (see also Fig. 2-5 in Chapter II for details)
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10 Supplementary information: Chapter III
Table S10-1 Comparison of geometric current densities for different strains of Shewanellaceae.
Strain jmax-CA/ µA cm-2
Applied E/ V vs Ag/agCl **Ref.
S. putrefaciens NCTC 10695 2.20 ± 0.62* -0.1 This study
S. putrefaciens NCTC 10695 3.43 ± 0.81* 0.0 This study
S. putrefaciens NCTC 10695 5.31 ± 1.47* +0.1 This study
S. putrefaciens NCTC 10695 7.76 ± 1.44* +0.2 This study
S. putrefaciens NCTC 10695 9.08 ± 1.70* +0.3 This study
S. putrefaciens NCTC 10695 12.03 ± 2.37* +0.4 This study
S. putrefaciens W3-18-1 3.1 MFC at 10 Ω [1]
S. putrefaciens SR-21 0.62 MFC at 1000 Ω [2]
S. putrefaciens ATCC 8071 31.25 MFC at 300 Ω [3]
S. putrefaciens IR-1 0.8 MFC at 1000 Ω [4]
S. putrefaciens IR-1 0.013 MFC at 500 Ω [5]
S. putrefaciens IR-1 0.002 +0.1 [6]
S. oneidensis MR-1 7.9 +0.2 [7]
S. oneidensis MR-1 5.1 +0.4 [8]
S. oneidensis MR-1 2.9 0 [9]
S. oneidensis MR-1 2.0 +0.2 [10]
S. oneidensis MR-1 45.0 +0.2 [11]
S. oneidensis MR-1 1.3 MFC at 10 Ω [12]
S. oneidensis MR-1 18.5 +0.043 [13]
S. oneidensis MR-1 10.0 -0.195 [14]
S. oneidensis MR-1 9.7 -0.195 [15]
S. oneidensis MR-1 17.8 +0.041 [16]
S. oneidensis MR-1 16.0 +0.041 [17]
S. oneidensis MR-1 22.5 +0.041 [18]
S. oneidensis MR-1 22.9 +0.5 [19]
S. oneidensis MR-1 23.6 +0.35 [20]
S. oneidensis MR-1 24.3 +0.2 [21]
S. oneidensis MR-1 25.7 0 [22]
S. oneidensis MR-1 9.6 MFC at 10 Ω [23]
S. oneidensis MR-1 IR-1< j < SR-21 MFC at 1000 Ω [24]
*Average data from chronoamperometric experiments at different applied potentials (vs. Ag/
AgCl) calculated as described in 3.2.4. and its respective standard deviation. **References in
Table: 1: (Bretschger, et al., 2010a); 2: (Kim, et al., 2002); 3: (Park and Zeikus, 2002); 4: (Kim,
et al., 2002); 5: (Kim, et al., 1999d); 6: (Kim, et al., 1999c); 7: (Carmona-Martínez, et al., 2011);
8: (Babauta, et al., 2011); 9: (Babauta, et al., 2011); 10: (Okamoto, et al., 2011); 11:
(Rosenbaum, et al., 2010a); 12: (Bretschger, et al., 2010a); 13: (Coursolle, et al., 2010); 14:
(Peng, et al., 2010b); 15: (Peng, et al., 2010a); 16: (Baron, et al., 2009); 17: (Marsili, et al.,
2008a); 18: (Marsili, et al., 2008a); 19: (Cho and Ellington, 2007); 20: (Cho and Ellington,
2007); 21: (Cho and Ellington, 2007); 22: (Cho and Ellington, 2007); 23: (Gorby, et al., 2006);
24: (Kim, et al., 2002).
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Table S10-1 Comparison of geometric current densities for different strains of Shewanellaceae
(…continuation of Table S10-1).
Strain jmax-CA/ µA cm-2
Applied E/ V vs Ag/agCl *Ref.
S. loihica PV-4 1.5 +0.2 [1]
S. loihica PV-4 4.0 +0.2 [2]
S. loihica PV-4 0.7 MFC at 10 Ω [3]
S. loihica PV-4 100 +0.2 [4]
S. loihica PV-4 1.0 +0.2 [5]
S. loihica PV-4 4.0 +0.2 [6]
S. loihica PV-4 6.0 -0.2 [7]
S. loihica PV-4 0.7 +0.2 [8]
S. loihica PV-4 1.6 +0.201 [9]
S. decolorationis NTOU1 34.0 +0.4 [10]
S. decolorationis NTOU1 97.0 +0.4 [11]
S. decolorationis NTOU1 22 MFC at 800 Ω [12]
S. japonica KMM 3299 22 MFC at 100 kΩ [13]
*References in Table: 1: (Wu, et al., 2011); 2: (Wu, et al., 2011); 3: (Bretschger, et al., 2010a);
4: (Zhao, et al., 2010a); 5: (Zhao, et al., 2010a); 6: (Liu, et al., 2010a); 7: (Liu, et al., 2010a); 8:
(Nakamura, et al., 2009b); 9: (Okamoto, et al., 2009); 10: (Li, et al., 2010); 11: (Li, et al., 2010);
12: (Yang, et al., 2011); 13: (Biffinger, et al., 2011).
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Table S10-2 Shewanella strains used as comparison in Table S10-1 and a description of their
isolation environment.
Strain Environmental characteristics of isolation area *Ref.
S. putrefaciens NCTC 10695 Oil emulsion from a machine shop
[1]
S. putrefaciens ATCC 8071 Responsible for butter putrefaction
[2]
S. putrefaciens W3-18-1 Marine sediment (630 m) in the Pacific Ocean,
Washington Coast, USA
[3]
S. putrefaciens IR-1 Anaerobic habitat in rice paddy field, South Korea
[4]
S. oneidensis MR-1 Anaerobic fresh water sediment in Lake Oneida, NY,
USA
[5]
S. amazonensis SB2B Shallow marine sediment (1 m) in the Amazon River
Delta, Brazil
[6]
S. putrefaciens SR-21 A transposon mutant of MR-1 with loss of Fe(III) and
Mn(IV) reduction
[7]
S. decolorationis NTOU1 Cooling system in an oil refinery in Taiwan
[8]
S. loihica PV-4 Microbial mat located at a hydrothermal vent in South
Rift of Loihi Seamount, Hawaii
[9]
S. oneidensis MR-4 Sea water, oxic zone (5 m) in the Black Sea
[10]
S. japonica KMM 3299 Sea water samples collected from a depth of 0.5-1.5 m
in the Gulf of Peter the Great, Sea of Japan
[11]
Note: Table partially made from information found in (Bretschger, et al., 2010a). *References in
Table: 1: (Pivnick, 1955); 2: (Derby and Hammer, 1931); 3: (Murray, et al., 2001); 4: (Hyun, et
al., 1999); 5: (Myers and Nealson, 1988); 6: (Venkateswaran, et al., 1998); 7: (Beliaev and
Saffarini, 1998); 8: (Chen, et al., 2008); 9: (Gao, et al., 2006); 10: (Nealson, et al., 1991); 11:
(Ivanova, et al., 2001).
Table S10-3 Cathodic and anodic peak positions, formal potential (vs. Ag/AgCl) and width of
potential window, ΔE, at a scan rate of 1 mV s-1
after SOAS baseline correction.
Applied E/ V Epc
/ mV Epa
/ mV Ef/ mV ΔE/ mV
-0.1 - - - -
0.0 - - - -
+0.1 - -7 ± 1 - 151 ± 12
+0.2 -129 ± 14 -6 ± 17 -67 ± 15 132 ± 20
+0.3 -126 ± 14 -1 ± 17 -64 ± 15 128 ± 23
+0.4 -118 ± 14 4 ± 16 -57 ± 15 127 ± 22
Epc
: cathodic peak position; Epa
: anodic peak position and Ef: formal potential. Window width
calculated according to Ref. (Firer-Sherwood, et al., 2008a).
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Figure S10-1 Electrochemical cell set-up. A) Electrochemical cell hosting six potentiostatic
controlled working electrodes without S. putrefaciens cells. B) Electrochemical cell with M1
growth media inoculated with whole cells of S. putrefaciens. Insert: photograph showing a
reddish pellet of S. putrefaciens formed when media was spinned down.
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Figure S10-2 Representative cyclic voltammograms for Shewanella putrefaciens biofilms
grown in the presence of (non-basal, e.g. 0.1 μM) higher levels of Riboflavin (1 μM).
Respective first Derivatives of each voltammogram are also shown, scan rate 1 mV s-1
.
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Figure S10-3 Effect of the Riboflavin concentration in the extracellular electron transfer.
Representative cyclic voltammogram of a Shewanella putrefaciens biofilm grown at a poised
(+0.4 vs Ag/AgCl) graphite electrode. The basal concentration of Riboflavin in the growth
media was 0.1 μM as reported in the Materials and Methods section (left panel). The
voltammogram was recorded at maximum biofilm activity after the start of the
chronoamperometry with a scan rate of 1 mV s-1
. Voltammetry of all Shewanella biofilms
grown at different applied potentials with no additional supplementation of Riboflavin (0.1 μM)
showed only one inflection point centered at 0 V (vs Ag/AgCl). After six semi batch
chronoamperometric cycles a pulse of fresh substrate containing 1 μM of Riboflavin was
injected into the electrochemical cell (right panel). For the experiment with additional
Riboflavin (1 μM) not only the inflection point at 0 V was observed but also an inflection point
centered at -0.4 V characteristic of the mediator molecule Riboflavin (Peng, et al., 2010b),
indicating that this molecule participated in the extracellular electron transfer process.
Furthermore, from the pronounced sharp rise of the inflection point centered at the midpoint
potential of Riboflavin, provided an example of how this mediator molecule increases the
electron transfer (Marsili, et al., 2008a).
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11 Supplementary information: Chapter VII
11.1 Influence of the buffer capacity
Figure S11-1 Influence of the buffer capacity: Cyclic voltammogramms (1mV s-1
) at pH 7,
wastewater derived and acetate–fed biofilms at varying buffer concentration, A) non-turn over
B) turn over conditions.
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11.2 Terminal restriction fragment polymorphism (T-RFLP) analysis: Anode biofilm
composition at the different pH values determined by T-RFLP
Figure S11-2 T-RFLP chromatograms (restriction digestion with RsaI and MspI) of the anode
biofilms formed at pH 7. The x axis represents the length of terminal restriction fragments and
the y axis the relative fluorescence units. On the right the area of every peak is shown as
percentage of the total peak area. The RsaI peak at 238 bp (503 bp with MspI) is shown in bright
yellow and represents Geobacter sulfurreducens (identified after sequencing).
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Figure S11-3 T-RFLP chromatograms (restriction digestion with RsaI and MspI) of the anode
biofilms formed at pH 9. The x axis represents the length of terminal restriction fragments and
the y axis the relative fluorescence units. On the right the area of every peak is shown as
percentage of the total peak area. The peak at 238 bp (503 bp with MspI) is shown in bright
yellow and represents Geobacter sulfurreducens (identified after sequencing). In the sample of
electrode-set 2 this organism could not be detected. This biofilm comprised several phylotypes.
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Figure S11-4 T-RFLP chromatograms (restriction digestion with RsaI and MspI) of the anode
biofilms formed at pH 6. The x axis represents the length of terminal restriction fragments and
the y axis the relative fluorescence units. On the right the area of every peak is shown as
percentage of the total peak area. The RsaI peak at 238 bp in the electrode-set 2 is shown in
bright yellow and represents Geobacter sulfurreducens (identified after sequencing the sample of
electrode-set 2). In the small dashed window the peak position is drawn to a larger scale to see
that the peak position of the RsaI peak is different in the sample of set 1 and set 2. The main
MspI peak is found at 161 bp that is also different from what was found for Geobacter
sulfurreducens in the other samples (Figures S11-2 and S11-3 above). This clearly shows that
Geobacter sulfurreducens could not be detected in the sample of electrode-set 1. This biofilm
comprised several phylotypes.
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Conclusion: T-RFLP showed a single peak after RsaI and MspI digestion which was affiliated to
Geobacter sulfurreducens after sequencing for both electrode sets grown at pH 7. The same
phylotype was also found at pH 6 and pH 9 but only in one (high performing) set, whereas in the
respective low performing electrode set no G. sulfurreducens could be detected.
11.3 Terminal restriction fragment polymorphism analysis: Anode chamber community
composition at pH 7 and 9 at different feeding cycles determined by T-RFLP
Figure S11-5 T-RFLP chromatograms (electrode-set 2, restriction digestion with RsaI) of the
replenished medium at the different feeding cycles. On the right the area of every peak is shown
as percentage of the total area. The peak at 238 bp is represented in bright yellow colour. It was
only found in samples of the feeding cycles at pH 7 and not in those at pH 9 (less than 1%). In
this figure, in comparison to the Fig. S11-2 above, a different resolution on the y axis was
chosen to give a better overview of the present diversity. Equal amounts of DNA were used for
the analysis of all samples.
11.4 Relationship of community composition when cultivated at different pH and under
successive feeding cycles determined by T-RFLP
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Figure S11-6 Similarity analysis derived from anode chamber communities when treated over
respective feeding cycles at pH 7 and 9 (all electrode set 2). As can be observed, the T-RFLP
derived composition of the pH 7 and 9 communities was clearly different. Undoubtedly, the
electrode biofilms were similar in T-RFLP composition for pH 6 and 7 whereas the biofilm
composition on the electrode treated at pH 9 was different (Analysis: non-metric MDS,
similarity measure: Bray-Curtis).
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11.5 Flow-cytometric analysis.
11.5.1 Community structure when cultivated at pH 9 at successive feeding cycles
determined by flow cytometry
Figure S11-7 Analysis of community structure by measuring the cells’ DNA contents and
Forward scatter behavior. Samples were harvested from the pH 9 anode chamber (electrode-set
2).
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11.5.2 Community structure when cultivated at pH 6 at successive feeding cycles
determined by flow cytometry
Figure S11-8 Analysis of community structure by measuring the cells’ DNA contents and
Forward scatter behavior. Samples were harvested from the pH 6 anode chamber (electrode set
2).
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11.6 Relationship of community structure when cultivated at different pH and under
successive feeding cycles determined by flow cytometry
Figure S11-9 Cluster dendrogram derived from anode chamber communities when treated over
several feeding cycles and at different pH. Feeding cycle numbers and pH affiliation are given
with c 1-5 and pH 6 to pH 9 (shown for electrode-set 2). As can be observed, the structure of the
inoculum community and that of the pH 9 electrode are clearly different from all other samples.
It is also obvious that distinct feeding cycles cluster together such as pH 7 c1 to c3, pH 6 c2 to
c4 and, pH 9 c2 to c4. It can be stated that similar micro-environments like successive feeding
cycles at a distinct pH value generated related community structures. A few of the pH related
communities clustered apart like pH 7 c4 to c5 or pH 6 c1 but are nevertheless close to each
other if the similarity analysis of Figure S11-9 is included. Undoubtedly, the electrode biofilms
were similar in structure for pH 6 and pH 7.
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11.7 Statistical Analysis of flow-cytometric data
For identifying community dynamics between feeding cycles and the different pH treatments a
newly developed method named ‘Dalmatian plot analysis’, a combination of image analysis and
multivariate community approach, was used (Bombach, et al., 2011). ‘Dalmatian plots’ are
simplified representations of the usually more complex flow cytometry bivariate plots.
Microbial sub-communities, detected by flow-cytometric measurements, are automatically or
manually encircled by black blots (‘gates’). Subsequently the underlying cytometry plot is
removed from the image. In a next step the resulting black and white Dalmatian plot is
converted into a binary (black-and-white), equal sized, pixel image having gate areas filled in
black and white as background.
For similarity calculation additionally estimates of overlapping areas from all possible binary
combinations are necessary. These are produced by overlaying all image combinations by
simple additive image calculation. For estimating the similarity rate between two images, the
area sums from all gates (the black area of a picture as pixels) from two images and the resulting
overlap of the same combination are estimated. Their similarity is then estimated using a
modified Jaccard index given by:
21
21
21
))2()1((1
AA
AA
AAOverlap
OverlapAAS
with similarity SA1A2 between two images A1 and A2, the sum of all gates in pixels counts of
picture A1 and A2, respectively and OverlapA1A2 the pixel sums of the overlap of image A1 and
A2. In this approach only presence and absence of sub-communities are regarded irrespective of
abundances within gates but thus enabling equal priority of all emerging sub-communities
therein.
Overlay creation and gate area calculation were automatically done with ImageJ Version 1.43
(http://rsb.info.nih.gov/ij). The similarity results of all possible combinations were then
transferred into a triangular similarity matrix. The final n-MDS analysis and a cluster analysis
based on the similarity matrix were accomplished under R Version 2.12.1 (Development-Core-
Team, 2010). For estimating overlays and estimating pixel areas of the single Dalmatian plots
and their overlays automatically a script for ImageJ was written.
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For estimating similarities and conduction of n-MDS and a cluster analysis directly from an
output of pixel counts created by ImageJ additionally a script for R was developed. Both scripts
are freely available and can be purchased by contacting the authors.
Figure S11-10 Illustration of methodology used for estimating community similarities of
cytometric flow plots using a Dalmatian-plot. Areas of gates were estimated as sum of pixels for
presence-absence when cell abundances taken into account. Sums were calculated from plots of
each of the samples separately and for the overlap of two samples, respectively. For similarity
estimation a modified Jaccard index was used (Figure S11-10 taken from (Müller, et al., 2011).
11.8 Biofilm detachment
Figure S11-11 Photograph of the detachment of a pH 7 grown biofilm from an electrode due to
extreme pH-conditions (pH 11).
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11.9 Multivariate statistical analysis of the flow-cytometric pattern using n-MDS-plots
The complex community and biofilm dynamics in response to micro-environmental changes like
pH-value and cycle conditions can be analyzed using Dalmatian plot based n-MDS analysis (for
method details see (Bombach, et al., 2011) and the following information presented in this
Chapter). In brief, first the microbial community changes detected via flow-cytometry are
visualized using Dalmatian-Plots. These plots are simplified representations of the more
complex density plots (see Figure 7-4), i.e. the raw-data, in which every dot represents a signal
event in the cytometric measurement. Within the derived Dalmatian plots black areas represent
identified microbial sub-communities (see below for an example and further explanation). Thus,
based on the number and position of these areas, representing sub-communities, every
Dalmatian plot can be considered as “fingerprint” of the bacterial culture for a certain point of
time and condition. Subsequently, based on these Dalmatian fingerprints, a similarity alignment
of the flow-cytometric data can be performed, which results in the n-MDS plot (Figure 7-5).
Roughly, within this plot, which is commonly used for the analyses of complex data sets, similar
samples are grouped together whereas dissimilar ones are grouped more distant. The stress-
value thereby provides a control measure for assessing the chance of data-misinterpretation. The
obtained stress-value for our analyses of 17.75 thereby clearly shows that the grouping of the
samples leads to no misinterpretation (see Figure 7-5).
-148-
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Curriculum Vitae
-A-
Alessandro A. Carmona-Martínez
Hagenring 30
38106 Braunschweig
Germany
Born August 19th 1982 Oaxaca, Mexico.
Telephone work: +49 531-391-8429
Mobile: +49 176-8604-6409
E-mail: [email protected], [email protected]
Career Objective
To seek a position in an academic or research institution that encourages innovative research and
promotes intellectual growth by utilizing my expertise and skills.
Education
2008-2012: Ph.D. (Dr. rer. nat.) in Biotechnology candidate (early 2012)
Institute of Environmental and Sustainable Chemistry. Group of Sustainable
Chemistry and Energy Research. Technical University of Braunschweig. Germany.
Thesis: “Study of the extracellular electron transfer processes between Shewanella
strains and electrode materials in bioelectrochemical systems”.
2005-2007: Master of Science in Environmental Biotechnology
Department of Biotechnology and Bioengineering. Centre for Research and Advanced
Studies of the National Polytechnic Institute. Mexico.
Thesis: “Electricity production in a microbial fuel cell fed with spent organic extracts
from hydrogenogenic fermentation of organic solid wastes”
2001-2005: Bachelor of Science in Environmental Engineering.
Interdisciplinary Professional Unit of Biotechnology of the National Polytechnic
Institute. Mexico.
Thesis: “Batch bio-hydrogen production with inhibited methanogenic consortia from
organic solid waste: effect of incubation temperature”
1997-2000: High school education, graduated in Mathematics and Sciences.
Curriculum Vitae
-B-
Peer-reviewed publications
1. A.A. Carmona-Martinez, K.H. Ly, P. Hildebrandt, U. Schröder, F. Harnisch*, D. Millo*,
Spectroelectrochemical analysis of intact microbial biofilms of Shewanella species for
sustainable energy production, In preparation, (2012).
2. A.A. Carmona-Martinez, F. Harnisch, U. Kuhlicke, T.R. Neu, U. Schröder, Electron transfer
and biofilm formation of Shewanella putrefaciens as function of anode potential,
Bioelectrochemistry, Accepted (2012).
3. S.A. Patil, F. Harnisch, C. Koch, T. Hübschmann, I. Fetzer, A.A. Carmona-Martínez, S.
Müller, U. Schröder, Electroactive mixed culture derived biofilms in microbial
bioelectrochemical systems: The role of pH on biofilm formation, performance and
composition, Bioresource Technology, 102 (2011) 9683-9690.
4. S. Chen, H. Hou, F. Harnisch, S.A. Patil, A.A. Carmona-Martinez, S. Agarwal, Y. Zhang, S.
Sinha-Ray, A.L. Yarin, A. Greiner, U. Schröder, Electrospun and solution blown three-
dimensional carbon fiber nonwovens for application as electrodes in microbial fuel cells,
Energy and Environmental Science, 4 (2011) 1417-1421.
5. S. Chen, G. He, A.A. Carmona-Martinez, S. Agarwal, A. Greiner, H. Hou, U. Schröder,
Electrospun carbon fiber mat with layered architecture for anode in microbial fuel cells,
Electrochemistry Communications, 13 (2011) 1026-1029.
6. A.A. Carmona-Martinez, F. Harnisch, L.A. Fitzgerald, J.C. Biffinger, B.R. Ringeisen, U.
Schröder, Cyclic voltammetric analysis of the electron transfer of Shewanella oneidensis MR-
1 and nanofilament and cytochrome knock-out mutants, Bioelectrochemistry, 81 (2011) 74-80.
7. H.M. Poggi-Varaldo, A. Carmona-Martínez, A.L. Vázquez-Larios, O. Solorza-Feria, Effect
of inoculum type on the performance of a microbial fuel cell fed with spent organic extracts
from hydrogenogenic fermentation of organic solid wastes, Journal of New Materials for
Electrochemical Systems, 12 (2009) 49-54.
8. I. Valdez-Vazquez, E. Ríos-Leal, K.M. Muñoz-Páez, A. Carmona-Martínez, H.M. Poggi-
Varaldo, Effect of inhibition treatment, type of inocula, and incubation temperature on batch
H2 production from organic solid waste, Biotechnology and Bioengineering, 95 (2006) 342-
349.
9. I. Valdez-Vazquez, E. Ríos-Leal, A. Carmona-Martínez, K.M. Muñoz-Páez, H.M. Poggi-
Varaldo, Improvement of biohydrogen production from solid wastes by intermittent venting
and gas flushing of batch reactors headspace, Environmental Science and Technology, 40
(2006) 3409-3415.
Curriculum Vitae
-C-
Oral/poster presentations in international conferences
1. A. Carmona-Martínez, S. Patil, F. Harnisch, U. Schröder, S. Chen, C. Greiner, A. Agarwal,
H. Hou, Y. Zhang, S. Sinha-Ray, A. Yarin. 2011. High Surface Area Electrospun and Solution-
blown Carbonized Nonwovens to Enhance the Current Density in Bioelectrochemical Systems
(BES). Abstract ELE 026. Presented at Wissenschaftsforum Chemie 2011, Bremen (Germany),
September 4th – 7th, 2011.
2. A. Carmona-Martínez, F. Harnisch, U. Schröder. 2010. Analysis of the electron transfer and
current production of Shewanella oneidensis MR-1 wild-type and derived mutants. Abstract P058.
Presented at Electrochemistry 2010: From microscopic understanding to global impact, Ruhr-
Universität Bochum (Germany), September 13th – 15th, 2010.
3. A. Carmona-Martínez, F. Harnisch, U. Schröder. 2009. Cyclic voltammetry as a useful
technique to characterize electrochemically active microorganisms: Shewanella putrefaciens.
Abstract AE15. Presented at Wissenschaftsforum Chemie 2009, Frankfurt am Main (Germany),
August 30th – September 2nd, 2009. ISBN: 978-3-936028-59-1.
4. A.A. Carmona-Martínez, 2009. Microbial fuel cells: an alternative for the production of
clean electricity. Abstract F128. Presented at German Academic Exchange Service Scholarship
Holders Meeting. Hanover (Germany). June 19th – 21th, 2009.
5. A. Carmona-Martinez, O. Solorza-Feria, H. M. Poggi-Varaldo. 2008. Batch tests of a
microbial fuel cell for electricity generation from spent organic extracts from hydrogenogenic
fermentation of organic solid wastes**. Abstract 2894. Presented at Third International Meeting on
Environmental Biotechnology and Engineering. Palma de Mallorca (Spain). September 21st - 25th
2008. ISBN: 978-84-692-4948-2.
6. A. A. Carmona-Martínez, O. Solorza-Feria, H. M. Poggi-Varaldo. 2008. Design and
characterization of a microbial fuel cell for electricity production from leachates**. Paper S001.
Presented at Sixth International Conference on Remediation of Chlorinated and Recalcitrant
Compounds. Monterey, California (USA). May 19th – 22th 2008. ISBN: 1-57477-163-9.
7. A. A. Carmona-Martínez, F. Esparza-García, J. García-Mena, O. Solorza-Feria, H. M.
Poggi-Varaldo. 2006. Actualidad y perspectivas en celdas de combustible microbianas para la
obtención de energía eléctrica a partir de residuales. Paper 109. Presented at Second International
Meeting on Environmental Biotechnology and Engineering. Mexico City (Mexico). September
26th – 29th 2006. ISBN: 970-95106-0-6.
**Presented at the congress by Dr. Héctor M. Poggi-Varaldo in my behalf.
Curriculum Vitae
-D-
Work and Research Experience
2012-2013: Coupling hydrogen production by dark fermentation and microbial electrolysis in a
single anaerobic reactor. References: Dr. Nicolas Bernet and Dr. E. Trably at the
Laboratory of Environmental Biotechnology of the French National Institute for
Agricultural Research.
2012: In vivo study of outer membrane cytochromes embedded in aggregations of living
bacteria (i.e microbial biofilms) grown on electrodes by a combination of surface-
enhanced resonance Raman scattering spectroscopy and electrochemistry. Reference:
Dr. Diego Millo at the Chemistry department/ Vrije Universiteit Amsterdam, the
Netherlands.
2008-2011: Experience on Bioelectrochemical systems (BES) aspects such as the extracellular
electron transfer mechanisms between bacteria and electrode materials in microbial
biofilms, analysis of environmental conditions affecting the performing of BES, study
of diverse electrode materials to enhance the performance of microbial biofilms in
microbial fuel cell systems, etc. References: Prof. Dr. Uwe Schröder and Dr. Falk
Harnisch at the Institute of Environmental and Sustainable Chemistry/ Technische
Universität Braunschweig, Germany.
2003-2007: Renewable biofuels (H2 and Biogas) production trough feasible and environmentally
friendly biotechnological process: e.g., hydrogen production from inhibited
methanogenic consortia. Reference: Dr. Héctor M. Poggi-Varaldo from the
Environmental biotechnology laboratory at the Centro de Investigación y de Estudios
Avanzados del Instituto Politécnico Nacional (Cinvestav), Mexico.
2005-2007: Design, construction and characterization of microbial fuel cells. Reference: Dr. Omar
Solorza-Feria at the Hydrogen and fuel cells laboratory, Cinvestav.
2003-2007: Use of analytical techniques for the detection of biotechnological compounds trough
methodologies based on gas and liquid chromatography. Reference: Mrs. Elvira Ríos-
Leal at the Analytic chemistry in biotechnology, Cinvestav.
2006-2007: Experience in the use of molecular tools for genetic typification of the microbioma
and microbial diversity in environmental and biotechnological systems. Reference: Dr.
Jaime García-Mena at the Laboratory of environmental genomics, Cinvestav.
Curriculum Vitae
-E-
Awards and Honours
2012: Foundation Caesar grant for post-doctoral research at the VU Amsterdam.
2008-2011: Ph. D. scholarship by the German Academic Exchange Service (DAAD).
2008-2011: Ph. D. complementary scholarship program by the Secretariat of Public Education of
Mexico (SEP).
2008: Winner in the student paper competition at the Sixth International Conference on
Remediation of Chlorinated & Recalcitrant Compounds (Monterey, California, USA)
2006: Winner of the poster competition in the renewable energies area at the Second
International Meeting on Environmental Biotechnology and Engineering (Mexico
City, Mexico).
2005-2007: M. Sc. scholarship by National Council of Science and Technology (CONACyT).
Professional memberships
-Mexican Talent Network e.V., Germany
-Mexican Society of Biotechnology and Bioengineering, A.C.
-Mexican Society of Hydrogen, A.C.
Linguistic skills Reading Writing Speaking
Spanish Mother tongue Mother tongue Mother tongue
English Proficient Proficient Proficient
German Good Good Good
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