29 Dipartimento di Fisica e Astronomia dell’Università and Sezione INFN,
Catania, Italy

30 Dipartimento di Fisica e Astronomia dell’Università and Sezione INFN,
Padova, Italy

31 Dipartimento di Fisica ‘E.R. Caianiello’ dell’Università and Gruppo
Collegato INFN, Salerno, Italy

32 Dipartimento di Scienze e Innovazione Tecnologica dell’Università del
Piemonte Orientale and Gruppo Collegato INFN, Alessandria, Italy

33 Dipartimento Interateneo di Fisica ‘M. Merlin’ and Sezione INFN, Bari,
Italy

34 Division of Experimental High Energy Physics, University of Lund, Lund,
Sweden

35 Eberhard Karls Universität Tübingen, Tübingen, Germany

36 European Organization for Nuclear Research (CERN), Geneva, Switzerland

37 Excellence Cluster Universe, Technische Universität München, Munich,
Germany

38 Faculty of Engineering, Bergen University College, Bergen, Norway

39 Faculty of Mathematics, Physics and Informatics, Comenius University,
Bratislava, Slovakia

40 Faculty of Nuclear Sciences and Physical Engineering, Czech Technical
University in Prague, Prague, Czech Republic

41 Faculty of Science, P.J. Šafárik University, Košice, Slovakia

42 Faculty of Technology, Buskerud and Vestfold University College, Vestfold,
Norway

43 Frankfurt Institute for Advanced Studies, Johann Wolfgang Goethe-
Universität Frankfurt, Frankfurt, Germany

44 Gangneung-Wonju National University, Gangneung, South Korea

45 Gauhati University, Department of Physics, Guwahati, India

46 Helsinki Institute of Physics (HIP), Helsinki, Finland

47 Hiroshima University, Hiroshima, Japan

48 Indian Institute of Technology Bombay (IIT), Mumbai, India

49 Indian Institute of Technology Indore, Indore (IITI), India

50 Inha University, Incheon, South Korea

51 Institut de Physique Nucléaire d’Orsay (IPNO), Université Paris-Sud, CNRS-
IN2P3, Orsay, France

52 Institut für Informatik, Johann Wolfgang Goethe-Universität Frankfurt,
Frankfurt, Germany

53 Institut für Kernphysik, Johann Wolfgang Goethe-Universität Frankfurt,
Frankfurt, Germany

54 Institut für Kernphysik, Westfälische Wilhelms-Universität Münster,
Münster, Germany

55 Institut Pluridisciplinaire Hubert Curien (IPHC), Université de Strasbourg,
CNRS-IN2P3, Strasbourg, France

56 Institute for Nuclear Research, Academy of Sciences, Moscow, Russia

57 Institute for Subatomic Physics of Utrecht University, Utrecht, Netherlands

58 Institute for Theoretical and Experimental Physics, Moscow, Russia

59 Institute of Experimental Physics, Slovak Academy of Sciences, Košice,
Slovakia

60 Institute of Physics, Academy of Sciences of the Czech Republic, Prague,
Czech Republic

61 Institute of Physics, Bhubaneswar, India

62 Institute of Space Science (ISS), Bucharest, Romania

63 Instituto de Ciencias Nucleares, Universidad Nacional Autónoma de México,
Mexico City, Mexico

64 Instituto de Física, Universidad Nacional Autónoma de México, Mexico City,
Mexico

65 iThemba LABS, National Research Foundation, Somerset West, South Africa

66 Joint Institute for Nuclear Research (JINR), Dubna, Russia

67 Konkuk University, Seoul, South Korea

68 Korea Institute of Science and Technology Information, Daejeon, South Korea

69 KTO Karatay University, Konya, Turkey

70 Laboratoire de Physique Corpusculaire (LPC), Clermont Université,
Université Blaise Pascal, CNRS–IN2P3, Clermont-Ferrand, France

71 Laboratoire de Physique Subatomique et de Cosmologie, Université Grenoble-
Alpes, CNRS-IN2P3, Grenoble, France

72 Laboratori Nazionali di Frascati, INFN, Frascati, Italy

73 Laboratori Nazionali di Legnaro, INFN, Legnaro, Italy

74 Lawrence Berkeley National Laboratory, Berkeley, California, United States

75 Moscow Engineering Physics Institute, Moscow, Russia

76 Nagasaki Institute of Applied Science, Nagasaki, Japan

77 National Centre for Nuclear Studies, Warsaw, Poland

78 National Institute for Physics and Nuclear Engineering, Bucharest, Romania

79 National Institute of Science Education and Research, Bhubaneswar, India

80 National Research Centre Kurchatov Institute, Moscow, Russia

81 Niels Bohr Institute, University of Copenhagen, Copenhagen, Denmark

82 Nikhef, Nationaal instituut voor subatomaire fysica, Amsterdam, Netherlands

83 Nuclear Physics Group, STFC Daresbury Laboratory, Daresbury, United Kingdom

84 Nuclear Physics Institute, Academy of Sciences of the Czech Republic, Řež u
Prahy, Czech Republic

85 Oak Ridge National Laboratory, Oak Ridge, Tennessee, United States

86 Petersburg Nuclear Physics Institute, Gatchina, Russia

87 Physics Department, Creighton University, Omaha, Nebraska, United States

88 Physics Department, Panjab University, Chandigarh, India

89 Physics Department, University of Athens, Athens, Greece

90 Physics Department, University of Cape Town, Cape Town, South Africa

91 Physics Department, University of Jammu, Jammu, India

92 Physics Department, University of Rajasthan, Jaipur, India

93 Physik Department, Technische Universität München, Munich, Germany

94 Physikalisches Institut, Ruprecht-Karls-Universität Heidelberg, Heidelberg,
Germany

95 Purdue University, West Lafayette, Indiana, United States

96 Pusan National University, Pusan, South Korea

97 Research Division and ExtreMe Matter Institute EMMI, GSI Helmholtzzentrum
für Schwerionenforschung, Darmstadt, Germany

98 Rudjer Bošković Institute, Zagreb, Croatia

99 Russian Federal Nuclear Center (VNIIEF), Sarov, Russia

100 Saha Institute of Nuclear Physics, Kolkata, India

101 School of Physics and Astronomy, University of Birmingham, Birmingham,
United Kingdom

102 Sección Física, Departamento de Ciencias, Pontificia Universidad Católica
del Perú, Lima, Peru

103 Sezione INFN, Bari, Italy

104 Sezione INFN, Bologna, Italy

105 Sezione INFN, Cagliari, Italy

106 Sezione INFN, Catania, Italy

107 Sezione INFN, Padova, Italy

108 Sezione INFN, Rome, Italy

109 Sezione INFN, Trieste, Italy

110 Sezione INFN, Turin, Italy

111 SSC IHEP of NRC Kurchatov institute, Protvino, Russia

112 Stefan Meyer Institut für Subatomare Physik (SMI), Vienna, Austria

113 SUBATECH, Ecole des Mines de Nantes, Université de Nantes, CNRS-IN2P3,
Nantes, France

114 Suranaree University of Technology, Nakhon Ratchasima, Thailand

115 Technical University of Košice, Košice, Slovakia

116 Technical University of Split FESB, Split, Croatia

117 The Henryk Niewodniczanski Institute of Nuclear Physics, Polish Academy of
Sciences, Cracow, Poland

118 The University of Texas at Austin, Physics Department, Austin, Texas, USA

119 Universidad Autónoma de Sinaloa, Culiacán, Mexico

120 Universidade de São Paulo (USP), São Paulo, Brazil

121 Universidade Estadual de Campinas (UNICAMP), Campinas, Brazil

122 University of Houston, Houston, Texas, United States

123 University of Jyväskylä, Jyväskylä, Finland

124 University of Liverpool, Liverpool, United Kingdom

125 University of Tennessee, Knoxville, Tennessee, United States

126 University of the Witwatersrand, Johannesburg, South Africa

127 University of Tokyo, Tokyo, Japan

128 University of Tsukuba, Tsukuba, Japan

129 University of Zagreb, Zagreb, Croatia

130 Université de Lyon, Université Lyon 1, CNRS/IN2P3, IPN-Lyon, Villeurbanne,
France

131 V. Fock Institute for Physics, St. Petersburg State University, St.
Petersburg, Russia

132 Variable Energy Cyclotron Centre, Kolkata, India

133 Warsaw University of Technology, Warsaw, Poland

134 Wayne State University, Detroit, Michigan, United States

135 Wigner Research Centre for Physics, Hungarian Academy of Sciences,
Budapest, Hungary

136 Yale University, New Haven, Connecticut, United States

137 Yonsei University, Seoul, South Korea

138 Zentrum für Technologietransfer und Telekommunikation (ZTT),
Fachhochschule Worms, Worms, Germany


#  Porous translucent electrodes enhance current generation from
photosynthetic biofilms

Tobias Wenzel 1, Daniel Haertter 1,2, Paolo Bombelli 3, Christopher J. Howe
3*, Ullrich Steiner 4*

We tested the enhancement of electrical current generated from
photosynthetically active bacteria by use of electrodes with porosity on the
nano- and micrometer length-scale. For two cyanobacteria on structured indium-
tin-oxide electrodes, current generation was increased by two orders of
magnitude and the photo-response was substantially faster compared to non-
porous anodes. These properties highlight porosity as an important design
strategy for electrochemical bio-interfaces. The role of porosity on different
length scales was studied systematically which revealed that the main
performance enhancement was caused by the increased surface area of the
electrodes. More complex microstructured architectures which spanned biofilms
as translucent 3D scaffolds provided additional advantage in the presence of
microbial direct electron transfer (DET). The absence of a clear DET
contribution in both studied cyanobacteria, Synechocystis and Nostoc, raises
questions about the role of conductive cellular components previously found in
both organisms.

1 Cavendish Laboratory, Department of Physics, University of Cambridge,
Cambridge CB3 0HE, United Kingdom
2 III. Physikalisches Institut, Georg-August-Universit t, 37077 G ttingen,
Germany
3 Department of Biochemistry, University of Cambridge, Hopkins Building,
Cambridge CB2 1QW, United Kingdom
4 Adolphe Merkle Institute, Rue des Verdiers 4, 1700 Fribourg, Switzerland
$*$ Corresponding authors<EMAIL_ADDRESS><EMAIL_ADDRESS>
Several microorganisms are able to generate electrons that can be collected
and utilised in external circuits [1]. In such devices, bio-anodes are the
electrodes that collect electrons from the living bio-catalyst. Bio-anodes in
the best studied bioelectrochemical technology, microbial fuel cells, are
commonly carbon or metal based, and a large diversity of morphologies has been
used [2]. The electrode porosity usually has a strong effect on device
efficiency [2], but the associated change in volume, surface area, and
organism contact area can rarely be disentangled from the variation of
materials themselves that are used to achieve the different morphologies.
These complicated correlations currently limit the understanding of design
rules for electrochemical bio-interfaces. Furthermore, because of a lack of
transparency of most anodes, there has been little work on the benefits of
using porous electrodes in microbe-based devices that rely on light
absorption, referred to as ‘biophotovoltaics’, except for one study using
larger, eukaryotic, algal cells [3]. While photosynthetic microorganisms are
expected to operate with a quantum efficiency of five to ten percent
internally, electrode interfaces and microbial electron export pathways
currently limit device efficiencies to much lower values [1].

In this study we tested the effect of electrode porosity at different length
scales on the performance of bioelectrochemical devices. To achieve this goal,
we have compared three different electrode morphologies of the same
translucent material. Two photosynthetic microorganisms _Nostoc punctiforme_
and _Synechocystis_ sp. PCC 6803 were each placed on (i) a non-porous indium
tin oxide (ITO) electrode, (ii) a thick ‘nanoporous’ ITO nanoparticle film,
and (iii) a ‘microporous’ inverse-opal structure made from the same
nanoparticles, and their photocatalytic current generation was investigated.

Doped metal oxides are popular transparent electrode materials for a wide
range of electronic applications. ITO is one of the best performing
transparent electrode materials, as it has a large optical bandgap, making it
transparent to visible light, while the high levels of tin doping cause a
metal-like conductivity. ITO is commonly used as thin film (tens of nanometres
thick) in display applications and was shown to be biocompatible [4, 5, 6].
ITO can be structured using a templating approach and has previously been used
as porous glass in electrochemical studies of enzymes [7, 8]. In order to
distinguish between the porosity on different length scales, we define
nanoporosity as the presence of pores between sintered nanoparticles (10–100
nm), and microporosity as the pores created by microsphere templates (10–40
µm). The templated inverse-opal pores used in this study are unusually large
and the electrodes are unusually thick (80–140 µm) in order to accommodate a
sufficient number of microorganisms within the structure to absorb incoming
light.

The two porosity length scales were chosen to represent biologically relevant
regimes of electron transfer from microorganisms to the anode. The
extracellular electron export mechanisms in cyanobacteria are still unclear,
even for the model organism _Synechocystis_ [1, 9]. Research on microbial
electrochemical devices distinguishes between direct electron transfer (DET)
from microorganisms to electrodes, and mediated electron transfer (MET)
facilitated by electrochemically active molecules in solution. Nanopores are
not directly accessible by the relatively large microbes but the increased
surface area is available for electrochemical interactions with redox-
molecules in the aqueous electrolyte, which is relevant for MET pathways. Pore
sizes comparable to the cell size in microporous morphologies allow cells to
enter the electrode, thereby providing a considerable increase in direct
contact area between bacteria and electrode surface.

Figure 1: BPV energetic scheme. Energy levels inside photosynthetic organisms
and charge transfer to external electrodes of biophotovoltaic devices.

Biophotovoltaic devices generate electric energy though photo-electrons
generated by living photosynthetic organisms on the anode, and a cathodic
reaction taking place at a lower potential, here recombination to water. The
energy levels involved in this process are detailed in the schematic diagram
in Fig. 1. The starting point of photosynthesis is photocatalytic water
splitting, during which the electrons are gained. This is achieved through the
absorption of two photons by the two photosystems (PSII and PSI), which form
the characteristic Z-Scheme. The derived electrons may be stored through the
production of NADPH molecules and can leave the cells via extracellular
electron transfer (MET & DET). The literature suggests similar values for
redox-potentials of electron export in known organisms [10] but with different
efficiencies [11]. Taking interfacial energy transfer losses into account,
these potentials determine the maximum voltage that can be generated in the
device, versus the fixed potential of a platinum air-cathode. In practice, the
cathode potential of air-cathodes does not reach the theoretical level of
water splitting [10], which is indicated by the higher position of the cathode
level in the scheme.

We used two different cyanobacteria in this study, _Nostoc punctiforme_ and
the model species _Synechocystis_ sp. PCC 6803. With either of these
microorganisms as bio-catalyst, both porous bio-anodes exhibited a remarkable
ca. 300 fold increase in generated peak photocurrent compared to non-porous
ITO films that are routinely used for biophotovoltaic devices. We also studied
the effects of the different anodes on the non-photosynthetic microorganisms
_Shewanella oneidensis_ and found similarly dramatic increases in external
current generation, with an additional enhancement on microporous electrodes.

## Results

Figure 2: Electrode porosity. (a) Scheme of microporous electrode fabrication
method. (b) Optical bright field microscopy image of an empty microporous
electrode (scale bar: 100 µm). (c) Optical dark field microscopy image showing
an electrode filled with cyanobacteria (after rinsing with water; scale bar:
20 µm). Scanning electron microscopy (SEM) images showing (d) the cross-
section of the electrode (scale bar: 40 µm) and (e) the nano-porous structure
of sintered nanoparticles (scale bar: 400 nm).

### Electrode design and characterisation.

A porous electrode that can incorporate photo-active biofilms must combine the
three qualities (i) conductivity, (ii) translucency and (iii) micro-porosity
on a length scale that allows cells to enter interconnected pores while also
forming biofilms with a thickness up to millimetres. To this end, an inverse
opal structure was designed with pores and pore-connections of several
micrometers, similar to that of cyanobacterial cells. The inverse opal
structure was generated through a templating approach (Fig. 2a). Polymer
micro-spheres of 40 µm in diameter were deposited to form an opal structure,
which was annealed to promote sphere adhesion and to control the diameter of
the sphere-sphere interconnects, followed by infiltration by a ITO
nanoparticle suspension via capillary forces. Filling the template with
nanoparticles ($<100$ nm) proved to be the fastest and most reliable method to
obtain thick porous films. A final heating step sintered the nanoparticles
while burning out the polymer template to leave the inverse structure behind.

The resulting material was diffuse white-yellow, as seen by eye or optical
microscopy (Fig. 2b). After the addition of cyanobacterial cells, the
electrode was examined by light microscopy (Fig. 2c), to confirm the ability
of the cells to populate the structure across the entire thickness (here, ca.
0.12 mm). Cells (ca. 2 µm cell-diameter) had to pass through ca. 10 µm wide
connections in order to reach all hollow spheres of the inverse opal
morphology (see Fig. 2b and cross-section in d) or reach these spaces via
cracks. The nano sized pores of the electrode material (Fig. 2e) could not be
accessed by the bacteria.

To compare the effects of micro-, nano- and no porosity in this study, three
different ITO structures were used. A commercial non-porous ITO layer on a PET
substrate served as reference (Fig. S1a,b), and thick nanoparticle films (Fig.
2e, Fig. S1c-f) without and with additional micropores (Fig. 2d, Fig. S1g,h)
were employed to assess the interplay of nano- and microporosity on the charge
generation by the microorganisms.

The nanoparticle film had a thickness of ca. 9 µm and was produced with the
same nanoparticles and on the same conducting FTO-glass substrate as the
microporous electrodes. Its sheet resistivity on a non-conducting substrate
was ca. 100 $\Omega$/cm2, determined by a 4-point probe. The film resistivity
was determined by impedance spectroscopy (see Methods). The resistivity of dry
electrode-‘sandwich’-samples with an area of approximately 0.25 cm2 were
measured as a function of film thickness. The obtained values were then
compared to that of thicker inverse opal structured electrodes, the
conductivity of which may be limited by their 3D morphology. The electrodes
exhibited purely ohmic resistance of 65 and 95 $\Omega$ for temperature
annealed ITO glass and 10 µm thick sandwiched nanoparticle films, respectively
(Fig. S2). The ca. 140 µm thick microporous electrodes displayed similarly low
purely ohmic resistance of ca. 115 $\Omega$.

The visual appearance and optical transmission spectra of the microporous and
the nanoparticle film are shown in Fig. S3. Ca. 50 % of the visible spectrum
was diffusely transmitted through the dry microporous film, and ca. 80 %
through the dry nanoparticle film. Non-porous ITO is fully transparent to
visible light, but causes partial specular reflection at air or water
interfaces due to its high refractive index. Nanoporous ITO (Fig. 2e) contains
many internal interfaces that scatter the light, rendering electrodes
translucent, which is even stronger for the inverse opal structure (Fig. 2d).
The relatively large opal unit cell size of 40 µm was chosen to minimise the
number of metal oxide water interfaces and thus the back-scattering of light,
and to allow the use of larger microorganisms. Scattering and absorption were
more strongly pronounced in this electrode than in previously described ITO-
based inverse-opal structures (e.g. in [12] ca. 80 % of the visible spectrum
is transmitted), due to the unusually large thickness of the electrode.

Figure 3: Design of biophotovoltaic devices. (a) Assembled and connected
3D-printed devices, incl. reference electrodes with with blue connectors,
positioned in the homogeneous illumination area. (b) Two schematic
perspectives of cuts through a device indicating the locations of electrodes
and biofilm.

### Biophotovoltaic devices.

Figure 3 pictures the electrochemical devices used for photo electricity
measurements in up to 12 parallel channels. The devices were 3D printed to
allow for a compact, shareable design and to enable temperature control via
hollow walls and a circulating water bath. The structured anodes on FTO-glass
were inserted from the bottom on a printed anode holder, and sealed with an
O-ring. The devices were designed to leave a (macroscopic) electrode area of 1
cm2 exposed to microorganisms and light. Three pieces of platinum nanoparticle
based air cathodes with a combined area of ca. 6 cm2 were arranged upright and
close to each anode opening. Parallel measurements with the same batch of
bacteria culture and under the same high-intensity light source were essential
to obtain reliable mean results in experiments.

Figure 4: Current levels by anode type and device configuration. (a)
_Synechocystis_ peak photocurrents in device mode (at ‘short-circuit’) and
half-cell operation (with reference electrode), during a photon flux of 460
µm/m2/s. (b) As (a), number of collected charges during 90 minutes of
illumination at photon flux of 512 µm/m2/s. (c) Current generated by
Shewanella cells for each anode type. (d,e) Schematic illustrating the device
and half-cell operation, respectively. The potentiostat connections for
reference (RE), working (WE) and counter electrode (CE) are indicated.

### Photocurrent ratios.

To study the effect of electrode porosity on the biophotovoltaic device
performance, parallel measurements with equal numbers of cells (based on
amount of chlorophyll) were conducted on the three different electrode
structures. The peak photocurrent levels observed with _Synechocystis_ are
shown in Fig. 4a. During the exposure of devices to light, the photo-current
did not stay constant, but reached a peak value, followed by a decreasing
slope, slowly reaching a steady value. The peak photocurrent was defined as
the average short-circuit current (0 V external potential between anode and
cathode and no reference electrode) measured during one minute around its peak
value, minus the dark-current level before each illumination, see Fig. S5.

Both micro- and nanoporous bio-anode based devices substantially outperformed
the non-porous ITO electrode with 11.5 and 8.4 mA/m2 to 0.04 mA/m2
respectively, while the microporous structure displayed a small but consistent
advantage over the film with nanopores only. During the illumination period of
seven minutes, the microporous devices reached a peak photocurrent value
$\sim$300 times higher than those equipped with non-porous electrodes.

In half-cell operation, the total charges collected during a longer period of
90 minutes were larger by more than a factor of 10 with microporous vs. non-
porous anodes, see Fig. 4b. Equilibrium values of the photocurrent are
difficult to obtain because BPV generated currents tend to fluctuate over
time. Photocurrent measurements are also limited by photo bleaching of
photosynthetic pigments during illumination and the long time periods
necessary for bio-anodes to stabilise after a change of conditions. To
quantify the photovoltaic output of the devices, the number of electrons
generated during 90 minutes of illumination (Fig. 4b) was therefore considered
as a robust current measure in addition to the recorded peak values (Fig. 4a).

Similar results were obtained for _Nostoc_ bio-anodes, where the current
enhancement of devices with a microporous electrode was also higher by a
factor of $\sim$300 compared to non-porous electrodes, with a peak-current of
11.2 mA/m2, and 30 times more charge collected during 90 minutes (167
µmol/m2).

Figure 4d illustrates the BPV device chamber at ‘short-circuit’ in which the
cathodic water recombination reaction drives the device current and voltage,
without an additional force of an externally applied potential.
Electrochemical studies commonly use a half-cell configuration instead (Fig.
4e). There, the anode potential is set with respect to a reference electrode
of a stable and known potential, here 0.2 V vs. an Ag/AgCl reference
electrode, and the cathode is dynamically shifted to a potential where the
cathodic reaction is non-limiting for the measurement. To test whether the
cathode limits the peak photocurrents reached in our devices, a set of peak
and continuum measurements were performed in half-cell mode, shown alongside
the device measurements in Fig. 4a,b. The highest current levels observed,
here in the case of porous anodes, were consistent between device and half-
cell operation within each others standard deviations, indicating the absence
of cathode limitation.

For comparison with an organism of well-studied ability to perform direct
electron transfer [13], an additional test was performed with non-
photosynthetic _Shewanella oneidensis_ bacteria in the three electrode set-up.
The average generated currents were 0.5 mA/m2, 62 mA/m2, and 299 mA/m2 for
non-porous, nanoporous, and microporous bio-anodes, respectively (Fig. 4c),
recorded in half-cell mode at an anode potential of 0.2 V vs. a Ag/AgCl
reference electrode in fresh LB medium. This corresponded to a more than
100-fold current increase from non-porous to nanoporous electrodes, and a
further ca. 5 times increase for the microporous structure.

Figure 5: Light response dynamics. Development of the average BPV photocurrent
over time at a photon flux density of 512 µmol/m2/s, for non-porous and
nanoporous film electrodes and both _Nostoc_ and _Synechocystis_.

### Light response characteristics.

Exposing photosynthetic electro-active biofilms to light gave rise to an
initial current peak which typically dropped to steady-state values. For each
of the organisms studied here, the photocurrent rose much faster to its peak
value on devices employing one of the porous compared to non-porous
electrodes, with little difference between the two pore types (Fig. 5 and S6).
The current peaks were reached for porous electrodes after 1–6 minutes,
whereas devices with non-porous electrodes required up to over one hour of
light exposure to reach a maximum (Fig. 5 and Fig. S6 expanded regions). The
steady-state current reached by non-porous bio-anodes was still lower compared
to the porous electrodes. Note that the photo-current minimum for
_Synechocystis_ cells on a non-porous electrode (Fig. 5) is unexpected. It was
not observed when the measurement was carried out with reference electrode
(Fig. S6). Since this study is concerned with the current maximum which was
reached after two hours, the origin of the minimum was not further
investigated.

Figure 6: Saturation behaviour. (a) Peak current generated by _Synechocystis_
cells on the three anode types at different light levels. The values are
averages of the peak minute after subtracting the dark current. (b) _Nostoc_
biofilm peak photocurrents as in (a).

The fast photo response of the porous bio-anodes enabled serial experiments
studying the peak BPV photocurrent for different light irradiances, see Fig.
6a,b. The resulting curves show an approximately exponential saturation of
photoelectron generation, with higher efficiencies (ratio of photocurrent to
photon flux) reached at low light levels. The performance advantage of porous
electrodes persisted for all irradiation levels. The similarity of
photocurrents on porous electrode types when used with either _Synechocystis_
or _Nostoc_ indicates the importance of the effective mesoscopic electrode
surface area rather than its coarse morphology. At the same time, the similar
saturation curves for each of these morphologies confirms the expectation that
the microporous electrode architecture does not shade cells to a significant
degree.

In addition to the use of porous bio-anodes, the need for fast and accurate
measurements was addressed (i) by adding phosphate buffer to the BG11 medium
which increased the electrolyte conductivity, (ii) by disregarding preceding
dark-current levels, and (iii) by choosing dark-times between (short)
illuminations that were long enough for the dark current level to recover. For
the low light level _Nostoc_ measurements (photon flux 0–400 µmol/m2/s, Fig.
6b), the irradiation intervals were not long enough to reach the full peak
values, which may explain the lower exponential slope for low photon fluxes
compared to the _Synechocystis_ data.

Surprisingly, in contrast to porous electrodes, non-porous ITO electrodes
performed better and responded faster in the absence of phosphate buffer, see
Fig. S7, but its performance remained much below that of the microporous
electrodes, which collected ten times more electrons during a 20 minute
illumination period.

### Redox reactions at the bio-anode.

The activity of electron-generation by bio-catalytic water splitting depends
only on the input energy of the illuminating light source (Fig. 1). By
measuring the electron collection as a function of anode potential by cyclic
voltammetry (CV) the redox activity of downstream electron donating molecules
can be characterised [14]. The bio-catalytic activity is slow for most
organisms, however, and could not be detected clearly by CV even at scan rates
as low as 0.5 mV/s. The porous anodes showed a strong enhancement in
electrochemical sensitivity, but also stronger surface charging, leading to
high charging currents (non-Faradaic) that broaden the CV hysteresis curve and
thus mask bio-catalytic peaks, see Figs. S8 and S9. This charge accumulation
is also reflected by the transient spikes observed in chronoamperometry
measurements when the light is turned off (e.g. Figs. 5 and S5).

Figure 7: Cyclic voltammetry analysis of microporous ITO anodes at a scan rate
of 4 mV/s with BG11 medium as electrolyte. (a) Response of the bio-anode
(_Synechocystis_) under exposure to different constant light levels. Light
intensity 1 corresponds to a photon flux of 820 µm/m2/s, and the time
indicates for how long the light level was applied by the end of the
measurement. (b) Scans of abiotic anodes in the dark. The oxygen content of
the electrolyte was varied by purging the electrolyte with nitrogen gas or
compressed air.

The increased electrochemical sensitivity of microporous electrodes did
however enable the detection of a light dependent reversible peak of a
reduction reaction on the anode surface (Fig. 7a). This peak was not observed
on the less sensitive non-porous ITO anodes. The slight shift in peak position
with irradiance was probably caused by a pH shift of the BG11-medium
electrolyte (no additional phosphate buffer was used in this measurement).
Lowering the oxygen content of the electrolyte by purging it with nitrogen gas
led to a decrease in peak height (Fig. S10), suggesting its link to
photosynthetic oxygen production. This was further confirmed by a measurement
without microorganisms, during which the oxygen content was varied by purging
with nitrogen or air, and for which the same reduction peak was reduced
(nitrogen) and enhanced (air), see Fig. 7b. The reaction appeared at anode
potentials lower than –300 mV, and was well separated from the operating range
of presented bio-anodes, which had a measured electrode potential of 50 to 150
mV vs. Ag/AgCl (Fig. 1). This separation is important for device performance,
as the electron-consuming reduction of oxygen can compete with the collection
of photosynthesis-derived electrons (oxidation reaction) if it takes place at
the material- and pH-dependent operating potential of the bio-anode.

## Discussion

Translucent conductive electrodes with porosity on two relevant length scales
– nanoporous films with pores accessible only by electrolyte, and microporous
electrodes with additional pores on the organism’s length scale – were
successfully created. Two cyanobacteria, _Synechocystis_ sp. PCC 6803 and
_Nostoc punctiforme_ , were applied to these electrodes where they continued
to be electroactive for the duration of measurements (at least several days),
with no indication of bleaching. During measurements, oxygen reduction at the
anode is unlikely to compete with the collection of microorganism-derived
electrons, as it takes place at lower anode potentials (–400 mV) than those
reached during BPV operation (ca. 100 mV).

In the architecture presented here, the cathode did not limit device
performance (observed peak photocurrents), and half-cell measurements were
equivalent to overall device performance. Measuring the bio-anode versus a
reference electrode (half-cell mode) simplifies the interpretation of results
of the anode performance and the presence of a reference potential allows the
application of methods such as cyclic voltammetry. However, half-cell
measurements can only be related to external quantum efficiencies as defined
in the field of solar energy generation, when it is known that an appropriate
non-limiting cathode such as the here-presented can be designed to complement
it in a full device.

Photocurrents collected from both types of porous electrodes were typically
one to two orders of magnitude larger than those from bio-anodes based on non-
porous ITO electrodes. The performance advantage varied depending on the
metric used (higher for peak currents; lower for charges collected over
several hours) and electrolyte conductivity (higher salt levels improved the
performance of porous anodes). Photocurrent peak values were also reached
faster (up to 90 times) on porous electrodes. The similar photocurrent levels
of both porous electrode types for each cyanobacteria species (Fig. 5)
indicates that the current enhancement can be attributed to a non-limiting
electrode surface area. The porous electrodes accordingly displayed similar
coulombic currents caused by surface-charging (Figs. S8 and S9). Both high
performance and fast response demonstrate the importance of a large electrode
surface area for the design of electrochemical bio-interfaces.

The fast response of porous bio-anodes enabled the performance of timely
serial experiments measuring the dependence of photocurrents on light
irradiation. The resulting curves show an approximately exponential saturation
behaviour. The correlation of this measurement to the saturated process of
photosynthetic electron transfer itself may point to a direct link between the
levels of photo-electrons available inside the cell, e.g. as the ratio of
NADPH/NADP+ molecules, and the peak number of photo-electrons measured in a
BPV device. The magnitude and temporal variation of photocurrent generation is
not yet well understood, and would depend on a complex interplay of
biochemical processes, including competition with the terminal oxidases, which
act as electron sinks [15]. Further research into the role of pigment
concentration, saturation of light absorption and the regulation of terminal
electron sinks within the cell will provide valuable insights into
biophotovoltaics, as well as into the bioenergetics of photosynthesis in
general. The quantitative datasets from the fast-response bio-anodes described
here could provide an important contribution towards this goal.

The high electrochemical sensitivity of the porous anodes, particularly the
microporous electrode enabling biofilm penetration, also enabled a sensitive
real-time detection of the oxygen levels present inside photosynthetically
active biofilms by cyclic voltammetry (Fig. 7). Such analyses might provide
quantitative insights into oxygenation studies e.g. in mixed biofilms or
tissues [16].

Surprisingly, the performance of nano- and micro-porous electrodes was similar
for both cyanobacteria. This similarity simplifies design rules of electrodes,
requiring simply the sintering of a nanoparticle paste for the manufacture of
nanoporous layers instead of more elaborate structuration. It also implies
that the dominating extracellular electron transfer mechanism may be self-
mediated (no additional electron shuttling molecules were added). If the
cyanobacteria were able to inject electrons efficiently into the electrode
surface via direct electron transfer (DET), a larger performance increase
would be expected for the microporous electrode, because it provides a many-
fold increase in organism-electrode contact area compared to the two films.
The potential DET-related performance increase was confirmed by using non-
photosynthetic bacterium _Shewanella oneidensis_ , which is well known to
exhibit DET [13] and for which microporous bio-anodes showed a clear (ca.
5-fold) additional advantage over nanoporous films. DET tends to be fast while
avoiding the diffusion losses associated with soluble electron carriers [10]
and can provide an important contribution to currents collected from
electroactive biofilms. So-called ‘nanowires’ have been proposed to give rise
to an efficient DET mechanism in some microbial biofilms. Nanowires are
conductive extracellular pili-like structures that transport redox-electrons
from within the cell through the insulating extracellular matrix and
neighbouring biofilm cells to electrodes [13, 17] but their electrochemical
properties are still disputed [18]. Conductive extracellular pili-like
structures (PLS), ca. 10 µm in length, have recently been identified in the
cyanobacteria utilised in this study, _Synechocystis_ sp. PCC 6803 and _Nostoc
punctiforme_ , as well as in the cyanobacterium _Microcystis aeruginosa_ , and
it was speculated that they contribute to extracellular electron transfer in
these organisms [13, 19, 20]. According to this hypothesis, the filaments
should lead to a strong enhancement in photocurrents when brought in contact
with the electrode surface across the entire photosynthetic biofilm, by using
the microporous structure. The performance similarity of biofilms on micro-
and nanoporous electrodes however suggests that the potential presence of
conductive bacterial filaments did not contribute significantly to the photo-
electron export in our devices, because the filaments did not connect to the
organism’s photosynthetic electron transfer chain or to the electrode, or the
filaments were not produced.

The measurements presented here suggest a slight advantage of the microporous
electrode morphology over the nanoporous film. This advantage might be due to
a larger, more accessible electrode surface, a smaller average distance self-
mediated small molecules have to travel to reach the electrode, or the fact
that biofilms reach further into the microporous electrodes,which therefore
probe the photoresponse of bacteria that are exposed to high irradiation
levels and thus contain a larger number of available photo-electrons. On the
other hand, the performance of microporous electrodes may be negatively
affected by increased scattering of the inverse opal structure, which leads to
slightly increased reflection and decreased transmission, as well as an
increase in electrode resistance with increasing thickness of the electrode.

It is clear that use of porous translucent electrodes offers a dramatic
increase in the current density obtainable from photosynthetic microorganisms
in biophotovoltaic devices. Their rapid photo-response times may additionally
allow these devices to be exploited as a wider tool for the study of
photosynthetic electron transfer.

## Methods

Making of inverse opal ITO electrodes. The microporous electrodes were
templated by polystyrene microshperes with an average diameter of 40 µm
(Dynoseeds TS 40 from Microbeads). For each electrode, 450 µl of a 5 %wt
suspension in deionised water were dried on a conductive FTO glass substrate
(8 Ohm/sq, Sigma Aldrich). A homogeneous area (ca. 2 cm2) of closed-packed
spheres was achieved when drying the suspension at 30$\,{}^{\circ}$C on the
substrate between two tightened aluminium sheets sandwiching a silicone
O-ring, and with a hole in the upper sheet of the same size as the O-ring
inner diameter. The frame can be used for convective assembly of colloids, but
the large microspheres here settled on the substrate quickly, decreasing the
influence of meniscal forces on the assembly. The bead structure was annealed
for 10 min at 130$\,{}^{\circ}$C on a hotplate. Nanoparticles were derived
from a commercial ITO nanoparticle dispersion ($<$100 nm) 30 %wt in
isopropanol (IPA) (Sigma Aldrich) by drying and re-suspending in absolute
ethanol (Sigma Aldrich) to result into a 10 %wt suspension. Ethanol was better
suited as a solvent because of its wetting properties with the polystyrene
opal. IPA leads to an increased coverage of the template top, leaving fewer
entry points for the microorganisms. The polystyrene opal was placed on a
hotplate at 45$\,{}^{\circ}$C for quick drying and filled with 3 times with 25
µl of ITO suspension. The template was burned out on a hotplate inside a fume
hood by heating the anodes to 500$\,{}^{\circ}$C, which simultaneously sinters
the nanoparticles, finishing the electrode. For this purpose, the following
heat ramp procedure was used: 3h heat ramp from room temperature to
300$\,{}^{\circ}$C holding the temperature for 1 minute; 10 min ramp to
325$\,{}^{\circ}$C holding for 5 min; 10 min ramp to 375$\,{}^{\circ}$C for 5
min; 10 min to 450$\,{}^{\circ}$C for 15 min; 10 min to 500$\,{}^{\circ}$C for
15 min; off. This method can be found in detail on DocuBricks.
For impedance spectroscopy testing, microporous electrode samples were
prepared on ITO-coated glass substrates (to avoid an additional material
interface) with the same protocol and annealed. They were then placed on
upside down on a second ITO-glass substrate which was freshly blade coated
with ITO nanoparticle paste (see below), and annealed again to sinter the
paste and provide a good contact. After the annealing, electrodes were filled
with room-temperature curing epoxy (EPO-TEK optical epoxy, 301-1LB kit) to
provide handling stability. Four samples containing 130-150 µm thick
microporous ITO layers made from the same nanoparticles had resistance values
of $110-165\,\Omega$.

Making of flat ITO electrodes. Plain ITO on PET electrodes with a surface
resistivity of 100 Ohm/sq. were used as purchased from Sigma Aldrich.
Nanoparticle film electrodes were made from the same commercial ITO
nanoparticle dispersion and FTO glass substrate as the microporous electrodes.
2.5 g equivalent of nanoparticles were mixed with 10.7 ml terpineol
(a-Terpineol 96+%, SAFC supply solutions). The IPA was evaporated off in a
rotary evaporator at 55$\,{}^{\circ}$C and vacuum pumping. FTO glass was taped
with kapton tape (Polyimide Tape from RS components) as distance spacer, and
the ITO-particle-terpineol paste was applied in between the tape strips.
Excess paste was removed by manually pushing the side of a glass pipette rod
over the spacers (blade-coating) to yield a plain film. The electrode was left
to settle at room temperature for ca. 20 minutes. The kapton tape was removed
manually. The electrodes were annealed with the same heat ramp protocol
described for microporous samples. This method can be found in detail on
DocuBricks. For impedance spectroscopy testing, ITO nanoparticle films were
prepared on commercial ITO-coated glass substrates (to avoid an additional
material interface) with the same blade coating protocol, and via spin-coating
at different speeds. Commercial ITO-coated glass samples without any
nanoparticle coating were annealed as the other samples, to provide an
appropriate reference point despite the deterioration of ITO conductivity
during the high temperature treatment. The mechanical junction of two
individually annealed film samples provided a less reliable contact than the
sintered microporous films, possibly causing a slight overestimation of their
ohmic resistivity.

Characterisation of porous electrodes. The transmission spectra in Fig. S3 of
the film were recorded with a Varian Cary 300 UV-Vis spectrophotometer, and
with an Ocean Optics FOIS-1 integrating sphere together with an Ocean Optics
USB4000 spectrometer; both using uncoated FTO-glass as optical reference
measurement (blank). The sheet resistivity of the nanoparticle film electrodes
was determined on a non-conductive glass substrate with four point probe (S302
Lucas Labs and Keithley 2400) to be 100 $\pm$ 10 Ohm*cm-2. The thickness of
the films was found to be 8.9 $\pm$ 0.6 µm by scanning over scratches with a
Dektak 6M stylus profiler. To probe the conductivity laterally, and through
the depth of the electrode film and compare these values to the microporous
inverse-opal structure, sandwich samples were built either by clipping two
nanoparticle film samples on ITO glass substrate on top of each other with two
paper-clips, or by sintering a microporous electrode onto a film (details in
respective electrode making method sections). The overlap area probing the
material films was about 0.25 cm2. The clean end pieces of the ITO-glass
substrates were contacted with silver paste and crocodile clips, and electrode
impedance spectroscopy measurements were performed across the enclosed films,
using the following parameters on a Biologic SP-300 Potentiostat: Scan at 0 V
from 1 MHz to 1 Hz, with 40 points per decade, a sinus amplitude of 5 mV,
waiting 0.1 periods before each frequency. No imaginary parts of the Nyquist
plot were found that could indicate a non-ohmic resistance behaviour of the
nanoporous or microporous films. The presented resistance data in this study
is thus simply an average of the (noisy but stable) real part of the
measurements.

Bacteria growth and quantification. _Synechocystis_ sp. PCC 6803 and _Nostoc
punctiforme_ cells were routinely cultured in BG11 medium supplemented with
about 1 mM NaHCO3 and maintained in sterile conditions at 30$\,{}^{\circ}$C
under continuous moderate light of 40-50 µmol photons m-2 s-1 and shaking at
160 rpm. Before applying cells to electrochemical devices, they were
concentrated via centrifugation (ca. 2000 g for 10 min). A small volume of
cells (120 - 160 µl) containing 134 nmol chlorophyll were pipetted into the
devices pre-filled with BG11 medium and after un-inoculated reference
measurements. The heavy cell suspension quickly settled onto the anode to form
a biofilm before the cell-suspension can mix with the bulk electrolyte. The
chlorophyll concentration was measured by extracting it from the cell
suspension in 99.8 % methanol (Sigma-Aldrich) and then calculating the
chlorophyll-a concentration from two optical densities, as described
previously [21].

Biophotovoltaic devices. The devices were printed on a Projet 3500HD Max 3D
printer with Visijet M3X acrylic material and an accuracy of 0.025 mm. The
support material (VisiJet S300) was washed out with hot sun-flower oil and
then IPA. Upright air-cathode windows (carbon paper with Pt-nanoparticles,
Alfa Aesar 45372 Hydrogen Electrode/Reformate) were sealed into the 3D printed
device with dental silicone (Zhermack, Elite HD+ Super Light Body). The anodes
were tightened in the device manually with screws and sealed with a nitrile
O-ring. A good contact between the anode edges and the connector-clamp was
ensured by conductive silver paste.

BPV operation and measurements. The BPV devices were loaded with the different
anode types and filled with a 10 ml volume of BG11 medium (containing small
amounts of phosphate) and phosphate buffer (DPBS 10x, D1408 Sigma-Aldrich, pH
7.2) in a mixing ration of 9:1.07 to obtain an overall phosphate buffer
concentration of 10 mM. Experiments referred to as ‘no added phosphate buffer’
used devices filled only with BG11 medium. The electrochemical measurements
were performed with a potentiostat (MultiEmStat by PalmSens) with 12
independent channels. The potentiostat applies a voltage and measures the
resulting current with a high resolution of down to 1 nA. Reference
measurements were conducted with the connected devices after at least 6 hour
waiting time (when chronoamperometry measurements seemed to have reached
equilibrium) and before adding cyanobacteria. Then, 134 nmol chlorophyll-a
equivalent of cyanobacteria were added to each device (see Bacteria growth and
quantification). Measurements were conducted again after a settling time of at
least 8 h and usually for a duration of one to three days. The illumination
periods with high light intensities were limited to five or seven minutes,
while more moderate photon flux densities of ca. 500 µmol/m2/s were used for
longer exposures. In the case of the custom LED white light source (Fig. S4),
the moderate illumination compares to the world average sun intensity reaching
ground level within the visible light spectrum. The temperature of the devices
was held stable at 25$\,{}^{\circ}$C by water cooling or heating of the hollow
devices with an external water bath.
The number of measurement channels per anode type used to obtain the presented
averages and standard deviations vary slightly because the devices that
shorted due to water condensation or leakage were not considered. During BPV
device measurements with _Synechocystis_ , all four devices each with micro-
and nanoporous electrodes and three devices with non-porous electrodes
delivered uninterrupted data. In half-cell mode, averages were formed of three
microporous electrode bearing devices, four with NP-films and three with non-
porous electrodes. For _Nostoc_ measurements, the numbers were two, four, and
two respectively. No increase in _Nostoc_ electro-activity was observed after
several days in the device, in contrast to the literature [20].
Measurements on _Shewanella oneidensis_ bacteria were conducted in half-cell
mode at an anode potential of 0.2 V vs. an Ag/AgCl reference electrode in
fresh LB medium without additional phosphate buffer. The same amount of
concentrated cell suspension was added to each device (1400 µl each, cells
were separated by repeated pipetting qith a 1 ml tip), and transmission
spectra were recorded at different cell densities as a point of reference for
cell numbers, see Fig. S11. Data was generated from four devices with micro-
and nanoporous electrodes each, and two non-porous ITO electrodes. The devices
were covered with Parafilm before measuring, which reduced the oxygen supply.
The Parafilm did not create fully anaerobic conditions, since during cyclic
voltammetry measurements, the presence of residual oxygen influx could still
be detected.

Microscopy. Optical microscopy was conducted with an Olympus BX60 microscope
and Olympus UMPlanFl 20x and 100x objectives. Images at manually adjusted
different focal depths were taken with an AxioCam MRc 5 (Zeiss) camera and
Z-stacked with CombineZP software. Slight adjustments were made to the colour
balance and contrast of the image. For electron microscopy images, a Leo
Gemini 1530 VP SEM was used with a Schottky-emitter consisting of a zirconium
oxide coated tungsten cathode and an in-lens secondary electron detector.

Data availability. The authors declare that the data supporting the findings
of this study will be made available in a public repository to which a
permanent reference will be included in the paper in time for publication.

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## Acknowledgements

We thank Antonio Abate for advice on conductivity testing across the films as
well as interpreting the impedance spectroscopy results and Erika Eiser for
discussion and the provision of colloids. TW is grateful for funding from the
Winton Fund for physics of sustainability, Mott Fund, Cambridge Trust (CHESS),
and the EPSRC CDT in Nanoscience and Nanotechnology. DH thanks Evangelisches
Studienwerk – Villigst e.V. for funding during this work. PB and CJH thank the
Leverhulme Trust for financial support. US acknowledges funding from the
Adolphe Merkle Foundation.

## Author contributions

TW, US, PB, and CH conceived the project and designed the experiments. TW
collected and analysed all included data and performed imaging. TW & DH
designed and built electrochemical devices, light source, and method for
microporous electrodes with advice from PB & US. TW wrote the manuscript,
further improved by all authors.

## Additional information

Competing interests: The authors declare no competing financial interests.

## Supporting Information for

Porous translucent electrodes enhance current generation from photosynthetic
biofilms

Tobias Wenzel, Daniel Haertter, Paolo Bombelli, Christopher Howe, Ullrich
Steiner

Figure S1: SEM images of the three anode types. (a,b) Commercial non-porous
ITO film on PET, with scale bars of 40 µm and 400 nm, respectively. (c)
Nanoporous film of sintered ITO nanoparticles, top view, with scale bar of 40
µm. (d,e,f) Cross-section of the nanoporous electrode, scale bars are 4 µm, 2
µm, and 400 nm, respectively; (e) also shows the underlying conducting FTO
layer on the glass substrate. (g,h) Top-view of a microporous electrode, with
scale bars of 40 µm and 500 nm respectively. The defects in (b) and (c) are
non-representative and are shown to visualise the smoothness of the
surrounding film. Figure S2: Resistance of different ITO nanoparticle film
thicknesses. Data points represent the ohmic resistance (averaged real part of
impedance measurement results) of dry nanoporous ITO ‘sandwich’ samples on an
ITO glass substrates, for different film thicknesses. Zero thickness refers to
a pair of ITO substrates without a sandwiched coating.  Figure S3: Macroscopic
appearance of the translucent electrode material. (a,b) Photographs of nano-
and microporous ITO nanoparticle films on FTO-glass substrates, respectively.
The microporous structure scatters visibly more light compared to the
nanoporous morphology. (b) Transmission spectra of the two electrode types in
(a) versus a FTO-glass reference. Integrating sphere measurements record the
total transmitted light, while the UV-VIS spectrometer data (dashed line) only
captures the light transmitted along the optical axis omitting the scattered
transmitted light. The microporous electrode transmits less light because of
increased scattering caused by the internal interfaces which leads to
increased backscattering. Figure S4: Illuminating light source. Spectrum of
custom LED light source (yellow) at maximum intensity compared to the solar
spectrum (red) (reference AM 1.5 Spectrum by ASTM, global tilt) at intensities
of 1000 W/m2 (solar simulator standard) and the world average sun intensity
reaching ground level (blue). Figure S5: Visual explanation of peak currents.
The first seven photocurrent peaks are shown of an illumination sweep with
increasing light intensity. The photocurrent peaks are indicated with stars
and numbers, along with data points during the peak minute. Averaging peak-
minute values provided a reliable value that was less dependent on noise and
peak shape. Illumination periods where chosen short enough (5-7 min.) and
separated enough (20-30 min.) to let the baseline dark-current recover before
the next illumination. To account for baseline fluctuations, the magnitude of
each photocurrent peak was measured from the preceding dark-current, here
indicated by red circles. The base level was also taken as average of data
during one minute. Underlying data is a chronoamperometry measurement fragment
of a _Synechocystis_ BPV with nanoporous electrode operated at short-circuit.
Figure S6: Light response characteristics of the three different
_Synechocystis_ bio-anode types over time, operated in half-cell mode at 0.2 V
vs. an Ag/AgCl reference electrode. The photon flux during illumination was
512 µm/m2/s. Averages and standard deviations (shaded) where formed from 3-4
individual devices per anode type. Figure S7: Current generated by
electrochemical devices (in half cell mode at 0.2 V vs. Ag/AgCl) with and
without biofilms (_Synechocystis_) operated at different light intensities.
The electrolyte in the devices was BG11 medium without supplementary phosphate
buffer. The temperature curve corresponds to the values measured in the
electrolyte close to the anode. In contrast to other measurements presented in
this study, the dark-times between illumination times were not chosen to be
long enough in this measurement for the dark current level to recover to its
pre-illumination base-level. Figure S8: Non-Faradaic charging current on
porous electrodes broadens cyclic voltammetry (CV) scans. Example CV data at
scan rate of 0.5 mV/s on _Synechocystis_ bioanodes differing in porosity is
shown. As indicated by arrows, data from scans performed on porous anodes are
broadened in comparison to non-porous electrodes (the later was ten times
enlarged for clarity) due to enhanced charging currents. They are also more
sensitive to detect electrochemically relevant molecules near the electrode,
as described in the results section. Figure S9: Data demonstrating the extend
of surface charging by anode type. Individual cyclic voltammetry scans were
taken at a scan rate of 0.5 mV/s with BG11 medium (without microorganisms and
light) on anodes differing in porosity. Charging currents were comparable for
both porous anode types but much lower for non-porous anodes. Figure S10:
Cyclic voltammetry analysis of a microporous bio-anode with _Synechocystis_
cells at a scan rate of 4 mV/s. Scans from -0.71 to 0.1 V vs. Ag/AgCl at a
photon flux density of 295 µm/m2/s. The tracks were recorded before, while and
after purging the electrolyte with nitrogen gas to remove the oxygen released
by the photosynthesising organisms. Figure S11: UV-VIS absorption spectra of
liquid Shewanella cell-culture at different dilution ratios. The amount of
concentrated cells between 5 µl and 50 µl, as indicated in legend, was added
to 1 ml of LB medium (reference solution) for each measurement. 140 µl of
concentrated cell-culture was used per device for electrochemical
measurements. This data serves as a reference for the amount of cells added,
in absence of an equivalent protocol for the determination of chlorophyll
concentration.