Investigation of electron transfer mechanisms in ...

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 Dissertation Kumulative Arbeit

von aus

Alessandro Alfredo Carmona Martínez Oaxaca / Mexiko

1. Referentin oder Referent: 2. Referentin oder Referent: eingereicht am: mündliche Prüfung (Disputation) am: Druckjahr 2012

Prof. Dr. Uwe Schröder Prof. Dr. Rainer Meckenstock 30.05.2012 05.10.2012

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. CarmonaMartínez, 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.

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 -i-

Table of contents (extended) 1

Extracellular electron transfer in Bioelectrochemical systems: bridge between

natural environments and applied technologies .......................................................................... 1 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

1.4.1.1

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

Microbial mediated extracellular electron transfer (MET) ................................................................ 8

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

Bioelectrochemical systems (BESs) .....................................................................................................11

1.5.1

Types of Bioelectrochemical systems ..............................................................................................13

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

1.6

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

Introduction ..........................................................................................................................................33

2.1.1

2.1.1.1

Direct electron transfer (DET) .................................................................................................34

2.1.1.2

Mediated electron transfer (MET) ...........................................................................................36

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

3

Extracellular electron transfer mechanisms of S. oneidensis MR-1 wild type and mutants .............34

Conclusions ..........................................................................................................................................46

Study of Shewanella putrefaciens biofilms grown at different applied potentials

using cyclic voltammetry and confocal laser scanning microscopy.. ................................. 47 3.1 3.1.1 3.2

Introduction ..........................................................................................................................................47 Influence of the electrode potential on electroactive microbial biofilms .........................................49 Materials and methods ..........................................................................................................................50

3.2.1


3.2.2


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


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 4.1

Introduction ..........................................................................................................................................61

4.2


4.2.1

Materials and methods .....................................................................................................................64

4.2.2


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


4.3.1 4.4

5

Bioelectrochemical current production ............................................................................................67 Conclusions ..........................................................................................................................................70

Electrospun and solution blown three-dimensional carbon fiber nonwovens for

application as electrodes in microbial fuel cells................................................................... 71 5.1

Introduction ..........................................................................................................................................71

5.2


5.2.1

Carbon fiber preparation ..................................................................................................................73

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

5.2.2

Electrode preparation .......................................................................................................................75

5.2.3

Bioelectrochemical experiments ......................................................................................................75

5.3


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 5.4

Cyclic voltammetry of electroactive biofilms grown at modified carbon electrodes .......................79 Conclusions ..........................................................................................................................................81

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6

Electrospun carbon fiber mat with layered architecture for anode in microbial fuel

cells...........................................................................................................................................82 6.1

Introduction ..........................................................................................................................................82

6.2


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


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

7

Conclusions ..........................................................................................................................................89

Electroactive mixed culture derived biofilms in microbial bioelectrochemical

systems: the role of pH on biofilm formation, performance and composition ................. 90 7.1

Introduction ..........................................................................................................................................90

7.2


7.2.1


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

7.2.8.1

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 7.3

T-RFLP and Sequencing .........................................................................................................95


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 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 8.3.1

Results and discussion ........................................................................................................................113 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

9

Conclusions ........................................................................................................................................118

Supplementary information: Chapter II .................................................................... 120

10 Supplementary information: Chapter III ................................................................... 130 11 Supplementary information: Chapter VII ................................................................. 136 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

12 References ...................................................................................................................... 148 -vi-

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 (Med red 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 redoxsystem 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 GESCFM 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

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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. ............................... 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, Red T 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, Red T 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 nonturnover 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

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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). ........................................... 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 TRFLP 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 pHconditions (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

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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.

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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.

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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).

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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 (Med red 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-

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 -5-

periplasmic decahaem c-Cyt MtrA, the outer membrane protein MtrB and the OM decahaem cCyts 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 coworkers (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-6-

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, -7-

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 (selfproduced 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

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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 selfproduced 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 Sphingomonas xenophaga Pseudomonas aeruginosa Pseudomonas chlororaphis Shewanella oneidensis Shewanella algae Bacillus pyocyaneus Propionibacterium freundenreichii Shewanella alga Acetobacterium woodii Pseudomonas stutzeri Methanosarcina thermophila Geobacter metallireducens Shewanella oneidensis Klebsiella pneumoniae a

Mediator molecule 4-amino-1,2-naphthoquinone Phenazine-1-carboxylic acid Phenazine-1-carboxamide Flavin mononucleotide Melanin

Reference (Keck, et al., 2002) (Price-Whelan, et al., 2006) (van Rij, et al., 2004) (von Canstein, et al., 2008) (Turick, et al., 2002)

Pyocyanine 2-Amino-3-carboxy-1,4naphthoquinone Cyanocobalamin Hydroxycobalamin Pyridine-2,6-bis Porphorinogen-type molecules Anthraquinone-2,6-disulfonate 1,4-Dihydroxy-2-naphthoate derivative Anthraquinone-2,6-disulfonate

(Friedheim and Michaelis, 1931) (Hernandez and Newman, 2001) (Workman, et al., 1997) (Hashsham and Freedman, 1999) (Lewis, et al., 2001) (Koons, et al., 2001) (Cervantes, et al., 2004) (Ward, et al., 2004) (Li, et al., 2009b)

More detailed information can be found in the following references: (Hernandez and Newman,

2001, Marsili and Zhang, 2010, Schröder, 2007, Watanabe, et al., 2009). -10-

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 halfcell 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). -13-

Table 1-2 Common terminology for the BES technology. Name Bioelectrochemical system

Abbrev. BES

Microbial fuel cell

MFC

Microbial electrolysis MEC cell

Definition An electrochemical system in which biocatalysts (microorganisms) perform oxidation and/ or reduction at electrodes A BES that produces net electrical power

Ref.* [1]

[2]

A BES to which net electrical power is provided to achieve [3] a certain process or product formation

Bioelectrochemically assisted microbial reactor Bio-electrical reactor

BEAMR A BES to which net electrical power is provided to achieve [4] a certain process or product formation

Biocatalyzed electrolysis cell

BEC

Biochemical fuel cell

BFC

Biofuel cell

BFC

Sediment microbial fuel cell

BER

A reactor in which current is provided to microorganisms [5] to stimulate their metabolism A BES to which net electrical power is provided to achieve [6] a certain process or product formation An electrochemical system in which biocatalysts function as catalysts for oxidation and/ or reduction reaction at electrodes An electrochemical system that use biocatalysts to convert chemical energy to electrical energy

[7]

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 An MFC for desalinating water based on using the electrical current generated by exoelectrogenic bacteria

[11]

An MFC that exploits the energy of light and the activity of phototrophic microorganisms to produce electricity

[13]

A new class of BES that uses whole organelles (e.g., mitochondria) as catalysts

[14]

Microbial desalination MDC cell Microbial solar cell

MSC

Mitochondrial biofuel MBFC cell

[8]

[12]

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 coworkers (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) hydrogengenerating 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 -16-

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 -18-

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 threedimensional 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 -19-

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 highthroughput 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 flowrate 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.

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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.

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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. -25-

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 ΔpilMQ/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). -26-

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.

16

A

A

A

14 +0.4 +0.3 +0.2 +0.1 0.0 -0.1

10

jmax/ A cm

-2

12

8 6 4 2 0

B

-2 0

1

2

B

B 3

4

5

6

7

8

time/ days 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.

-27-

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.

-28-

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.

-29-

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.

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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. -31-

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 setups 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 electrochemical 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.

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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 -33-

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.

-34-

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.

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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 knockout 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)). -36-

2.2.3 Bioelectrochemical experiments The

bioelectrochemical

experiments

were

carried

out

under

potentiostatic

control

(Potentiostat/Galvanostat VMP3, BioLogic Science Instruments, France) utilizing a threeelectrode 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 nonturnover 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 semibatch 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. jmax / µA cm-2 Strain

Mutant description

∆flg

Flagella deletion mutant

Wild Type

S. oneidensis MR-1 wild-type, ATCC 700550

ΔpilM-Q

Type IV pilus deletion mutant

∆mtrc/

Outer membrane decaheme

∆omcA

c-type cytochromes MtrC and

This

Ref.

Ref. Ref. Ref.

worka

Ab

Bc

Cd

De

9.5 ± 2.0

8

-

-

-

7.9 ± 1.5

5

13

3.4

6

7.7 ± 3.7

7.6

-

-

-

4.3 ± 0.8

0.6

2

0.7

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


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 -83-

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 (2DECFM) 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 JSM7500F equipment under a voltage of 5 kV. The energy dispersive X-ray analysis (EDX) was -84-

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


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 Porosity/ Resistivity/ Geometric current size/ μm % Ω cm 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 layeredCFM, 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 -85-

the oxidation of the substrate (acetate) catalyzed by microbial biofilms at the working electrode according to the following equation: CH 3COO   H 2 O

bioelectrocatalyse



2CO2  7 H   8e 

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. -86-

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 -87-

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. -88-

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. -90-

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


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. -91-

7.2.2 Electrochemical set-up All electrochemical experiments were carried out under potentiostatic control, using threenecked-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 pHvalue 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 -92-

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 -93-

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 -94-

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 longpass 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

(http://folk.uio.no/ohammer/past/).

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analysis

was

done

with

PAST

7.3


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 -96-

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 pHenvironments 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 nonturnover 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 -100-

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 pHgradient 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, flowcytometric 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 electrodeset 2, were investigated on their microbial structure (by flow-cytometry) and composition (by TRFLP) 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 flowcytometric analysis of a bacterial anode chamber community after wastewater inoculation. Five -101-

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 NebeVon-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 fedcycle 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 pHvalues 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 75). 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. -104-

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. 16), 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 -109-

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 -110-

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


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 threenecked-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 -112-

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


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 highcurrent-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. -114-

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 nonelectrogenic 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 electrochemical 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 nonchanging 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 nonturnover CVs (Harnisch, et al., 2011). -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. 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 followup 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 -118-

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 5.1 +0.4 2.9 0 2.0 +0.2 45.0 +0.2 18.5 +0.043 10.0 -0.195 9.7 -0.195 17.8 +0.041 16.0 +0.041 22.9 +0.5 23.6 +0.35 24.3 +0.2 25.7 0 * Average maximum current density

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This study (Babauta, et al., 2011) (Babauta, et al., 2011) (Okamoto, et al., 2011) (Rosenbaum, et al., 2010a) (Coursolle, et al., 2010) (Peng, et al., 2010b) (Peng, et al., 2010a) (Baron, et al., 2009) (Marsili, et al., 2008a) (Cho and Ellington, 2007) (Cho and Ellington, 2007) (Cho and Ellington, 2007) (Cho and Ellington, 2007)

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 nonturnover 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/ΔmshHQ. 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 S. putrefaciens NCTC 10695 Oil emulsion from a machine shop

*Ref. [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 Anaerobic habitat in rice paddy field, South Korea

[3]

Anaerobic fresh water sediment in Lake Oneida, NY, USA Shallow marine sediment (1 m) in the Amazon River Delta, Brazil A transposon mutant of MR-1 with loss of Fe(III) and Mn(IV) reduction Cooling system in an oil refinery in Taiwan

[5]

S. putrefaciens IR-1 S. oneidensis MR-1 S. amazonensis SB2B S. putrefaciens SR-21 S. decolorationis NTOU1 S. loihica PV-4 S. oneidensis MR-4

[4]

[6] [7] [8]

Microbial mat located at a hydrothermal vent in South [9] Rift of Loihi Seamount, Hawaii 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 [11] in the Gulf of Peter the Great, Sea of Japan 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: S A1 A 2  1 

( ( A1)   ( A2)   OverlapA1 A 2 )

 Overlap

A1 A 2

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-CoreTeam, 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 stressvalue 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).

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Curriculum Vitae 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.

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Curriculum Vitae 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 threedimensional 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 MR1 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. PoggiVaraldo, Effect of inhibition treatment, type of inocula, and incubation temperature on batch H2 production from organic solid waste, Biotechnology and Bioengineering, 95 (2006) 342349. 9. I. Valdez-Vazquez, E. Ríos-Leal, A. Carmona-Martínez, K.M. Muñoz-Páez, H.M. PoggiVaraldo, 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.

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Curriculum Vitae 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 Solutionblown 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, RuhrUniversität 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.

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Curriculum Vitae 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 surfaceenhanced 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 RíosLeal 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.

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Curriculum Vitae 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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