Environ. Sci. Technol. 2010, 44, 2721–2727
Quantification of Electron Transfer
Rates to a Solid Phase Electron
Acceptor through the Stages of
Biofilm Formation from Single Cells
to Multicellular Communities
J E F F R E Y S . M C L E A N , * ,† G R E G W A N G E R , †
YURI A. GORBY,† MARTIN WAINSTEIN,†
JEFF MCQUAID,† SHUN’ ICHI ISHII,†
ORIANNA BRETSCHGER,†
HALUK BEYENAL,‡ AND
K E N N E T H H . N E A L S O N †,§
The J. Craig Venter Institute, San Diego, CA, The Gene and
Linda Voiland, School of Chemical Engineering and
Bioengineering and Center for Environmental, Sediment and
Aquatic Research, Washington State University, Pullman, WA,
and University of Southern California, Los Angeles, CA
Received October 11, 2009. Revised manuscript received
February 8, 2010. Accepted February 13, 2010.
Microbial fuel cell (MFC) technology has enabled new
insights into the mechanisms of electron transfer from
dissimilatory metal reducing bacteria to a solid phase electron
acceptor. Using solid electrodes as electron acceptors
enables quantitative real-time measurements of electron
transfer rates to these surfaces. We describe here an optically
accessible, dual anode, continuous flow MFC that enables realtime microscopic imaging of anode populations as they
develop from single attached cells to a mature biofilms. We
used this system to characterize how differences in external
resistance affect cellular electron transfer rates on a per cell
basis and overall biofilm development in Shewanella
oneidensis strain MR-1. When a low external resistance (100
Ω) was used, estimates of current per cell reached a
maximum of 204 fA/cell (1.3 × 106 e- cell-1 sec-1), while
when a higher (1 MΩ) resistance was used, only 75 fA/cell
(0.4 × 106 e- cell-1 sec-1) was produced. The 1 MΩ anode
biomass consistently developed into a mature thick biofilm with
tower morphology (>50 µm thick), whereas only a thin
biofilm (<5 µm thick) was observed on the 100 Ω anode.
These data suggest a link between the ability of a surface to
accept electrons and biofilm structure development.
Introduction
Dissimilatory metal reducing bacteria (DMRB) can utilize
solid phase electron acceptors such as iron and manganese
oxides as electron acceptors to respire anaerobically via a
process termed extracellular electron transport (EET). This
ability has important implications for metal cycling in the
environment, the ecophysiology of these organisms, their
potential use for bioremediation (1–3) and also microbial
fuel cell applications (4, 5). Three different and perhaps
* Corresponding author e-mail: jmclean@jcvi.org.
†
The J. Craig Venter Institute.
‡
Washington State University.
§
University of Southern California.
10.1021/es903043p
 2010 American Chemical Society
Published on Web 03/03/2010
overlapping strategies for coordinating extracellular electron
transfer to solid phase electron acceptors are apparently used:
direct cell-mineral contact with multiheme outer membrane
cytochromes serving as reductases (6), small molecular weight
compounds that function as electron shuttles (7, 8), or
electrically conductive bacterial pili or nanowires (9, 10).
Because of their EET abilities, DMRB are commonly used
in microbial fuel cells (MFCs) (4, 5), where they are capable
of EET to MFC anodes as they anaerobically oxidize organic
matter. In particular, model organisms such as Shewanella
or Geobacter species have been used in efforts to unravel the
biological mechanisms involved with current production
(6, 9–14). In such studies it is especially critical to quantify
cell numbers as well as describe the depth and architectures
of biofilms as they develop on MFC electrodes. However,
achieving this knowledge is difficult given typical MFC system
designs. Commonly used designs to maximize power generation, cannot be used to accurately quantify the number
of cells per electrode surface area or the depth of the biofilm;
often this can only be achieved at the terminus of the
experiment, which does not provide a direct link between
cell colonization/growth and power production. Further, the
sampling methods chosen for such quantifications must be
delicately employed so that biomass or attachment is not
affected by mechanical disruption (15).
Destructive single end-point analyses such as total protein
(14), and qualitative imaging using various microscopic
techniques, render it impossible to study biofilm development, morphology, or architecture through time. Recent
mathematical models of electron transfer in biofilms (16)
indicate the importance of a conductive biofilm in the transfer
of electrons; therefore, biofilm parameters including depth
must be measured. Comparative studies of current generation
must be directly related to the number of cells on the electrode
surface and electron transfer rates. This is germane when
using mutant strains, which may be altered in attachment,
biofilm formation, and/or rates of EET. To determine the
rate of electron transfer per cell it is critical to first obtain an
accurate count of the number of cells on the anode. The
inability to accurately quantify attached cells and normalize
for them inhibits strict standardizations across samples,
organisms, and MFC designs among the greater MFC
community and even within individual laboratories.
One solution to this dilemma is to merge the technology
used to measure electron transfer to solid surfaces with the
tools for live, noninvasive imaging. A number of studies have
used optical flow cells and electrochemically controlled
electrode surfaces such as indium tin oxide (17) to investigate
how different potentials affect bacterial detachment (17),
attachment, and/or growth rates (18). Recently Franks et al.
(19) demonstrated the utility of a “mini-stack” MFC that was
modified for nondestructive visualization of biofilm growth,
a study that focused on changes of pH within anode biofilms.
We describe here the construction and application of a
parallel plate flow cell with incorporated fuel cell components
that accommodates real-time fluorescent microscopic imaging with real-time measurements of current production. With
this system, we report for the first time, the electron transfer
rates on a per cell basis for Shewanella oneidensis MR-1
interacting with anode electrodes; as well as the changes in
anode potential through the development stages of a biofilm
from single attached cells to a mature, multicellular, threedimensional structure.
VOL. 44, NO. 7, 2010 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
9
2721
Materials and Methods
Parallel Plate Flow Anode Chamber with Air Cathode. The
optically accessible MFC was a modified, commercially
available, biofilm flow cell (model no. FC-81; Biosurface
Technologies Inc.). These flow cells were modified by
removing the glass slide substrates and replacing them with
electrode materials. The anodes were made from superfine
isomolded graphite plates (Graphite Store) that were shaped
into 5 × 25 × 1 mm electrodes fitted with 0.25 mm platinum
wire leads and polished to a final mirror finish with 12000
grit polishing paper. Silicone sealant was used to cover all
surfaces of the anodes except the polished top, yielding a
total accessible anode surface area of 1.25 cm2. The aircathode consisted of Pt (5 g/m2) coated graphite felt (GDE
LT 120E-W, The Fuel Cell Store) coated with Nafion 117 film
(Fluka) that was painted on to the surface and allowed to
air-dry (20). By placing two anodes side by side in the same
compartment, we fit two MFCs into one chamber which
allowed us to compare their performances under identical
operating conditions. Similar design was used by Dewan et
al. (21), to compare power of many MFCs which had multiple
anodes. The two anodes were connected to the shared aircathode and cell voltage was measured individually across
100 Ω and 1 MΩ resistors to determine current under
operating conditions (Iop) and current density (i) with a digital
multimeter (Keithley 2700, Keithley Instruments Inc.). The
anode and cathode electrodes were separated from each other
by a nonconductive plastic slide. These MFCs were specifically designed to direct the growth medium across the anode
surfaces before flowing into the cathode compartment
(Supporting Information (SI) Figure 1). At the operating flow
rate of 200 µl/min the flow was calculated to be laminar
(Reynolds number of 4.3). The chamber volume was 300 µL
with a calculated average flow velocity of 16 mm/min
translating to a dilution rate of 0.67 min-1 or 1.5 volume
changes each minute.
Microorganism, Growth Medium, Buffer, and Electrolyte
Solution. Shewanella oneidensis strain MR-1 p519nGFP
(constitutively expressed green fluorescent protein) was used
for all experiments (22). For each experiment, cells from a
frozen stock were streaked onto Tryptic-Soy Agar (Difco)
plates augmented with Kanamycin and incubated for 18 h
at 30 °C. These pregrown cells were transferred to 20 mL of
modified M1 minimal medium (6) and incubated for 24 h at
30 °C and at 150 rpm. To ensure an excess of the electron
donor lactate in all experiments the M1 medium was
supplemented with 60 mM lactate. Measurement of the MFC
effluent by high pressure liquid chromatography (HPLC)
confirmed the excess of lactate (data not shown). To minimize
the concentration of oxygen in the system the influent
medium was continuously sparged with ultrahigh purity N2.
Fresh medium was delivered into the MFC using a variable
speed peristaltic pump (404, Ismatec).
Flow Cell Operation. The MFCs where chemically sterilized using a 10% peroxyacetic acid solution and flushed with
sterile MFC buffer containing 50 mM PIPES, 100 mM NaCl
at pH 7.2. Prior to inoculation, background measurements
were taken in MFC buffer solutions, with and without lactate,
as well as the minimal growth medium. Inoculation of the
MFCs involved injecting the cells immediately upstream of
the chamber and allowing them to flow into the chamber
and attach to the graphite anodes for 10 min with the flow
off (batch mode). Following this, a high flow rate (i.e., 10 min
at ∼2 mL/min) was set to flush out any unattached cells. The
flow rate was then decreased to the operational level of 200
µL/min.
Measurements and Calculations. Potentiodynamic polarizations were used to calculate power curves. These data
were used to determine the maximum current (Imax), maximum current density (imax), and maximum power densities
2722
9
ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 44, NO. 7, 2010
(Pmax) of MFCs ((5)) based on the true surface area of the
anode. The anode was the limiting electrode and cathodic
current was in excess. We used a potentiostat (Reference
600, Gamry Instruments, Inc.) and a three-electrode system
with the anode as the working electrode, the cathode as the
auxiliary electrode and Ag/AgCl as the reference electrode.
The anode potentials were scanned from the open circuit
cell voltage (OCV) to a cell potential of 0 mV using a step size
of 0.1 mV/s and a sampling time of 1 s. The cathode produced
a stable cathodic potential of +230 mVAg/AgCl throughout the
experimental period as was measured before and after cell
polarization. The resulting increase in cell potential observed
at open circuit indicates the anode potential becoming more
negative with respect to the cathode over time. A Student’s
t test was used for statistical comparisons of anode performance at 120 hrs, assuming a normal distribution and equal
variance. The internal resistance of the designed MFCs were
measured with active biofilms using the Electrochemical
Impedance Spectroscopy (EIS) method described by Dewan
et al. (21), and found to be 40 Ω.
Microscopy (Epifluorescent, Confocal, Scanning Electron). Epifluorescence. For real time imaging of the cells on
the anode surfaces a Zeiss Axio microscope with fluorescence
capabilities was used to image 10 random fields-of-view per
anode surface at each set time interval to determine the
average cell numbers. These counts were typically conducted
without disruption of the operation of the MFC or flow rates
except for a brief interruption during the removal of the MFC
from the microscope after the 18 h time point (Figure 1)
resulting in a discharge of anode potential through a short
circuit. Image processing was done by means of ImageJ and
the biofilm analysis package ISA 3D software (23). The effect
of fluorescent imaging on the MFC’s was also conducted
with and without cells showing no change in the cell voltage
due to any of the excitation wavelengths used.
Confocal Microscopy. Microscopic observations were
performed on a Leica TCSP5 confocal laser scanning
microscope (CLSM) (Leica Microsystems). Image stacks were
obtained using a 10× objective. Fluorescence for GFP was
collected with 488 nm laser emission. Simulated 3-D images
and sections (plan views) were generated using the software
Volocity and the plan views with side profile slices using
IMARIS (Bitplane AG, Z|Aaurich, CH).
Scanning Electron Microscopy. At the termination of each
experiment, anodes were fixed in a 2.5% solution of glutaraldehyde, processed through an ethanol dehydration series
(i.e., 25, 50, 75, 100% v/v Ethanol, 0.5 h each treatment), then
critically point dried (815 Auto-Samdri, Tousimis) (10). The
anodes were mounted on aluminum stubs, coated with
chromium and imaged in a Phillips XL30 Environmental SEM.
3.1. Results and Discussion
Variation in Biomass and Electrochemical Parameters with
External Resistance. Using the dual anode real-time imaging
compatible MFC we successfully and repeatedly grew biofilms
on the anode surface, monitored cell surface growth,
quantified biomass and calculated operational current (Iop)
with different external resistance. Experiments were very
reproducible, and we show here one representative data set
from one typical experiment with fluorescent images (Figure
1). The development of current over time followed a general
trend for all repeated experiments with a lag in current for
several hours followed by an exponential phase and a
relatively stable current phase. The corresponding image data,
showing how biomass changed during current production,
revealed a consistent and striking difference developing
between the biomass on the anodes. After 80 h, a thick and
highly structured biofilm had formed on the 1 MΩ anode
while the biofilm on the 100 Ω remained as a thin layer of
cells. Using noninvasive CLSM imaging, the biofilm structure
FIGURE 1. Simultaneous monitoring of current generation and biofilm development from single attached cells with epi-fluourescent
imaging. Temporal current changes in the imaging compatible MFC for two different anodes with external resistances of 1 MΩ (A)
and 100 Ω (B). C) Concurrent image data taken over the course of the experiment for both anodes. Differences in biofilm
development were clearly visible by 88 h. Scale bars equal to 10 µm for 0 and 15 h and 100 µm for 18-130 h.
FIGURE 2. Representative CLSM and SEM images of the anodes after 5 days of growth. CLSM images collected nondestructively
demonstrate the biomass and structural differences between the 1 M Ω (A) and 100 Ω (B) anodes. Representative SEM of the 1 M Ω
(C) and 100 Ω (D) processed by critical point drying confirm the structural differences in the biofilms as well as indicate that the 1
M Ω colonies contain a greater amount of extracellular material between the attached cells within these colonies.
and thickness were quantified for both anodes. Thicker
biofilms with qualitatively more extracellular material were
observed on 1 MΩ anode when compared to biofilms that
formed on the 100 Ω anode (Figure 2). The biofilms on the
100 Ω anodes appeared as low mounds of cells rising only
∼5 µm from the anode surface (Figure 2b) whereas the 1 MΩ
anodes produced a thick base layer with large “towers” (50-80
µm) (Figure 2a). This tower morphology is commonly
observed in conventional continuous flow biofilm growth
chambers where soluble compounds, such as oxygen or
fumarate, serve as the terminal electron acceptor (22, 24).
Images collected using SEM were consistent with those
obtained using CLSM, despite an observed loss of biofilm
biomass during critical point drying for SEM analysis. These
VOL. 44, NO. 7, 2010 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
9
2723
FIGURE 3. Representative timecourse measurements of
attached cell counts and current density during operation over
24 h on the 1 M Ω and 100 Ω anode. A) Average cell counts
per cm2 of anode area (left axis) and the current density during
operation (right axis). B) Timecourse plot of the current
generated per cell based on direct in situ cell counts and the
stable operating current.
data demonstrate that higher resistance (lower current flow)
maximizes biofilm development or alternatively a lower
resistance(highercurrentflow)minimizesbiofilmdevelopment.
Current and Cell Density Correlation. Current production and cell counts were monitored from initial attachment
through the early stages of biofilm formation on the anode.
Initial cell densities on the anodes averaged 2 × 105 cells/
cm2 with virtually no change in numbers during the first 15 h
of observation. During this initial period the anodes were
covered with only a single layer of cells but the current density
under operation increased during this time (Figure 3),
indicating an increase in specific anode respiration rate.
Between 15 and 18 h, a rapid increase in cell numbers and
current was detected for both anodes. By 24 h cell densities
reached 1.2 × 106 cells/cm2 with the 1 M Ω, and 8.8 × 105
cells/cm2 with the 100 Ω. Operational current density also
increased to 0.09 µA/cm2 (1 M Ω) and 0.08 µA/cm2 (100 Ω)
over this period.
Specific Electron Transfer Rates. Current generation was
calculated on a per cell basis using the electrochemical and
microscopic data obtained in real time (Figure 3). The
temporal trends in these experiments indicate that the current
per cell maximum during operation was reached before the
cells went into the rapid growth phase on the anode surface.
The 100 Ω anode in the first run (Figure 1) increased initially
to a maximum of 200 fA/cell (1.3 × 106 e- cell-1 sec-1) and
the 1 MΩ reached 80 fA/cell (0.4 × 106 e- cell-1 sec-1) (Figure
3). These rates decreased to 63 fA/cell and 39 fA/cell,
respectively, after the cell growth entered an exponential
phase. Replicate runs conducted over a longer time scale (75
h) (SI Figure 2) demonstrated that the trends in cell growth
and current generation were very consistent, although the
absolute values as well as the time required for rapid increases
in cell numbers and current densities showed some variability.
To our knowledge, these results represent the first time
the absolute values of current per cell over time have been
estimated from direct cell counts for a bacteria within an
operating MFC. These are also the first estimates of the
specific electron transfer rates to an anode surface for
Shewanella oneidensis strain MR-1. Previous studies have
estimated the current per cell from total protein concentra2724
9
ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 44, NO. 7, 2010
tions at selected time points during a batch operated MFC
for Geobacter sulfurreducens with values of 23 fmol/cell/day
(14). At the values obtained for MR-1 on the anode operating
with a 100 Ω resistor, we obtained ranges from 200 to 63
fA/cell which corresponds to specific electron transfer rates
of 179 to 56 fmol/cell/day during continuous operation mode.
As noted above, the average cell concentration on the
anodes during the first 15 h did not change significantly,
whereas the current increased in a linear fashion during this
initial period (Figure 3). To determine if, given enough time,
the initial cells bound to the surface would produce detectible
current without the components in the medium for growth
and division, the experiment was modified such that after
inoculation and for the following 48 h, MFC buffer augmented
with 60 mM lactate was substituted for the M1 growth
medium allowing respiration without growth. This experiment showed no observable increase in cell numbers or any
detectible increase in current during this time (data not
shown). After 48 h the MFC buffer was switched to the
standard growth supporting M1 minimal medium and the
current and cell numbers began to increase as described
above. This suggests that the increased current production
is dependent upon growth or perhaps newly synthesized
cellular components needed for extracellular electron transfer
such as redox active compounds, that is, more redox active
compounds per cell. While initial attachment on the two
anodes were in similar ranges, consistently more cells
attached to the 1 MΩ anode. At later time points, when
discrete cells were no longer resolvable on the anode surface,
the image analysis software package, ISA 3D (23), was used
to quantitatively compare the biomass differences and
determine the correlation of colony size (average diffusion
distance) with current development (SI Figure 3).
Temporal Development of Anode Potential and Maximum Current Density. Polarization measurements were
made at several times during the experiment, in an effort to
determine how the anode potential and maximum current
density (imax) correlated with biofilm development. For these
measurements (Figure 4), the system was placed in open
circuit for 2 h with uninterrupted flow, and the anode
potential monitored against an Ag/AgCl reference electrode
(see SI Figure 1) to ensure the anode potential had stabilized.
The overall shape of the anode potential vs time curve
(Figure 4) was observed to be similar between the two anodes.
However, the 1 MΩ anode established a lower anode potential
and produced a higher imax more rapidly than the 100 Ω
anode. The 100 Ω anode began to rapidly increase after 100 h
and reached nearly the same potential as the 1 M Ω. However,
neither polarization curve correlated with the timing of the
Iop increase trend and the period of exponential cell growth.
The anode potential therefore became negative (reducing)
and the imax increased in a linear fashion from 20 to 60 h with
only a monolayer of cells present on the anodes and did not
correlate to the rapid current development period (30 to 60 h).
At 20 h of growth with a single layer of cells and before rapid
cell growth, the anode potential reached -150 mVAg/AgCl,
consistent with the increasing current in the initial growth
stages as shown in Figure 3. From 60 to 120 h the Iop of the
MFC’s decreased slightly over time whereas the anode
potential continued to become more negative and the
imaxincreased.
The higher cell potential and performance of the MFC at
later times is consistent with reported literature for batch
studies (6). Interestingly, in the early stages of the experiment,
the potential and power densities are higher and are
established sooner in the 1 MΩ anode than the 100 Ω anode,
but by 120 h the 100 Ω anode approaches this value. Since
the 100 Ω anode did not support a thick biofilm by 120 h,
the development of its potential and Pmax may not necessary
be reliant on a thick biofilm suggesting that a limit on a
FIGURE 4. Development of anode potential and performance over the course of biofilm development. Current vs time for (A) 1 M Ω
and (B) 100 Ω anodes. The spikes in cell voltage observed in panels A and B represent the times where the fuel cell was in open
circuit and when the polarization data were collected. Results from polarization resistance from anodes in A and B over time; (C)
development of anode potential. (D) development of maximum current density.
MFC’s power is reached prior to the maximum cell density
achieved on an anode.
Comparing MFC Performance Parameters with Pronounced Biomass Differences. At 120h, when the biofilm
thickness differences between anodes were very pronounced,
potentiodynamic polarization measurements were performed for five separate experiments (two anodes within
each chamber), and the mean of the data were used to
quantify the differences in MFC performance with the
different external resistance (SI Figure 4). The stable operating
current established after 120 h showed the largest measurable
difference between the two anodes, even while suffering from
high variability in the performance of the 100 Ω anode. The
100 Ω anode statistically outperformed the 1 MΩ anode with
mean stable operating currents of 1.2 ( 0.6 µA and 0.3 ( 0.03
µA, respectively. The observed differences are consistent with
standard MFC theory that the maximum operating current,
or power, is achieved when the fuel cell is operated under
conditions where the internal and external resistances are
nearly balanced (e.g., 100 Ω external and ∼40 Ω internal in
this system) (5).
The open circuit potential of the anodes at this later time
point, although on average higher in the 1MΩ circuit, was
not significantly different (p ) 0.061) at the 5% level (p )
0.05), despite the clear difference in biofilm thickness (SI
Figure 4). The maximum current density however, was
significantly higher (p ) 0.027) on the 1 MΩ. The Pmax is on
average higher in the 1 MΩ circuit (4.5 ( 1.0 mW/m2) than
that of the 100 Ω (3.0 ( 0.9 mW/m2) though not significantly
so (p ) 0.084).
Overall, the current density and power density values fall
into a range of operating values for several reported microbial
fuel cells using Shewanella and show our system is operating
as well as other designs using similar electrode materials.
The most relevant comparison for a power measurement
that was believed to be predominately biofilm related was
reported by Biffinger et al. (25) at 0.52 mW/m2 for Shewanella
strain DSP10. They employed a Pt/C oxygen reduction
cathode and attempted to control for a strict biofilm
measurement with little influence of planktonic cells. The
power densities reported by Biffinger et al. are comparable
to those found in this study because both values were
calculated using the true surface area of the anode electrode,
and not the underestimated “projected” surface area of a
porous electrode, which can dramatically increase the
calculated power density (26).
Discussion
The optical compatible dual anode system described here
has allowed, for the first time, real time, noninvasive estimates
of current per cell collected noninvasively over time in an
operating MFC. In the initial phase, the 100 Ω anode increased
to a maximum of 200 fA/cell (1.3 × 106 e- cell-1 sec-1) and
the 1 MΩ reached 80 fA/cell (0.4 × 106 e- cell-1 sec-1). These
rates decreased to 63 fA/cell and 39 fA/cell respectively after
the cell growth entered a rapid phase of growth. The average
current on a per cell basis over time for both anodes in the
initial stages fell within a range of values from 0.007 to 0.2
pA per cell with the 100 Ω, which had a lower cell density
on the anode surface, outperforming the 1 M Ω. One
hypothesis as to why the current per cell decreases as the cell
surface growth enters exponential phase is that some
electrons are diverted to building cellular components rather
than to respiration via the anode ((5)).
As cells were observed to populate the anode surface, a
relatively consistent lag phase in current was seen before
rapid cell growth began and the stable current was attained.
One possible explanation for the observed lag phase in current
production could be that initially the cells are not adapted
to transferring electrons to solid phase electron acceptors
and that upregulation of genes encoding specific proteins
and production of new components for this transfer are
required. The cells from the liquid grown batch culture used
as the inoculum of the fuel cells are from the late exponential
phase and most likely the population is not homogeneous
and can even include metabolically inactive cells that still
maintain fluorescence from the long-lived GFP in this strain.
Our cell counts would include the dead or inactive cells that
still have intact membranes, which may or may not have
enzymatic activity and contribute to the overall current.
Bretschger et al. (6) however, have previously shown that
membrane fractions of cells alone do not produce current.
Overall there are difficulties associated with obtaining a true
value for the current per cell and therefore these data
represent estimates that are relevant to the specific conditions
of our system. Using a different fluorescent construct that
harbors a short-lived GFP however, may allow an estimate
of anode respiration rates for only the active cell population.
Previous studies using noninvasive NMR spectroscopy
on live biofilms of S. oneidensis MR-1 under flowing conditions in similar minimal medium, quantified the anaerobic
metabolism rates with lactate and various soluble electron
acceptors (i.e., fumarate, DMSO, NO3) (22) as well as
metabolite profiles within an MR-1 biofilm (27, 28). It was
VOL. 44, NO. 7, 2010 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
9
2725
found that even aerobically grown biofilms could immediately
reduce fumarate, DMSO or NO3 without prior exposure to
these acceptors or adjusting the oxygen concentrations to
anaerobic conditions. These reduction studies were conducted under bulk aerobic conditions where oxygen becomes
rapidly limited at depth within a metabolically active biofilm.
Here we have quantified the electron transfer to solid electron
acceptor and the changing potentials from single attached
cells though to mature biofilms. Quantification of parameters
such as the decreasing anode potential (becoming more
negative with respect to the reference electrode) over time
indicated that the cells can establish reducing conditions
early in the growth phase while still as a single cell layer. For
both the thick biofilms on the 1 MΩ anode as well as the thin
but active 100 Ω anode biofilm, the anode potential increased
rapidly when the circuit was opened, reaching reducing
conditions within 2 h. Presumably this is due to the cell’s
oxidation of lactate coupled to the reduction of all available
redox active compounds within the cells and possibly any
extracellular redox components. It is very interesting that
current was produced and anode potential was measurable
above background under constant flow with only a thin
dispersed monolayer on the anode. Given that this was under
relatively high laminar flow conditions and high dilution rates,
nonadhered soluble redox active compounds, such as
electron shuttles, would be of minimal use with single cells
on the surface. Of particular note is that the maximum rate
of electron transfer to the anode occurred at cell densities
that did not completely saturate the available anode surface
area. We suspect that a very low level of oxygen is also present
in solution since maturation of fluorescent proteins actually
requires the presence of molecular oxygen. Fluorescence
formation of GFP is prevented by rigorously anoxic conditions
(<0.75 µM O2) but is readily detected at 3 µM O2 ((29)). This
suggests that the cells (as a single cell layer) on the surface
are limited for oxygen as the acceptor and/or prefer the anode
as the electron acceptor; an intriguing idea that warrants
further investigation.
MFC performance parameters, determined from power
curves, were averaged for five independent runs. The 120 h
time point was chosen for the comparisons since that time
showed the most pronounced difference in biomass and
stable operating current (SI Figure 4) between the two anodes.
These results may be of use in determining whether thick or
thin biofilms are optimal for current generation in MFC
applications. The averages at 120 h show that the cell
potential, imax, and Pmax is higher in the 1MΩ circuit. However,
at 120 h, only the imax, was shown to be significantly higher
at the 5% level (p ) 0.05). Further study is needed to fully
characterize if these parameters are significantly increased
when a larger biofilm is present on the anode.
Although we have documented that varied external
resistance reproducibly results in major biofilm differences,
the explanation of these differences remains unknown.
Previous work with MR-1 biofilms in flow cells (on inert
surfaces) showed their development into complex structures
in nutrient rich (24, 30) and in a defined minimal medium
(22) grown with oxygen as the major electron acceptor. The
development and stability of these structures in MR-1 can
vary depending on the availability of a soluble alternative
electron acceptor such as fumarate (22). Noninvasive temporal development of MR-1 into biofilms from single attached
cells to mature biofilms has not been well documented or
previously assessed on an anode of a microbial fuel cell or
in correlation with electron transfer rates. It is possible that
the availability of an electron acceptor (potential difference
between the anode and cathode) or alternatively the rate at
which electrons can be passed to an accepting surface
(resistance) can drive the building of biomass. Increasing
the number of cells or redox active compounds to generate
2726
9
ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 44, NO. 7, 2010
a more reducing environment would be an effective strategy
to more quickly reach conditions to enable reduction of solid
phase oxides. The observed rapid biomass accumulation and
the more negative anode potential observed on the harder
to reduce (higher resistance) surface supports this notion.
In summary, we have successfully demonstrated a new
MFC design to couple imaging technology with microbial
fuel cells enabling insights into the physiology of microbial
cells as they develop into a biofilm especially as it pertains
to the movement of electrons from cells to a solid acceptor.
The analogy of these systems to the interaction of microbes
in the environment with solid mineral phases should not be
overlooked. Microbial fuel cells stand as a technology that
can be used to investigate the physiology of the cells on a
solid electron accepting surface and as a model system for
quantifying cell-mineral electron transfer. Several models
have been proposed detailing the mechanisms of electron
transport in MR-1 to solid metal oxides such as direct
attachment, electron shuttles and, the most recently discovered, microbial nanowires however the exact mechanisms
and the relative contributions of each are unclear. Further
studies with optical compatible systems may help decipher
the role of various mechanisms in solid phase electron
transfer. In particular the use of real-time fluorescent gene
reporters of the various redox proteins, pili, and electron
shuttles would help understand their temporal and spatial
(depth) related expression profiles.
Acknowledgments
This research was supported by the J. Craig Venter Institute;
the Legler-Benbough Foundation; and the Office of Science
(BER), US Department of Energy with funds from the
Environmental Remediation Science Program (grant DEFG02-08ER64560).
Appendix A
Nomenclature
Iop
i
imax
Pmax
current under operating conditions (A)
current density based on the true anode surface
area (A/m2)
maximum current density based on the true surface
area of the anode (A/m2)
maximum power density based on the true surface
area of the anode (watts/m2)
Supporting Information Available
Four figures intended to supplement the material presented
in the manuscript. Diagram of the optically compatible MFC,
replicate timecourse measurements for current per cell
estimates, biofilm structure and current relationships, and
5 day MFC performance parameter comparisons. This
material is available free of charge via the Internet at http://
pubs.acs.org.
Literature Cited
(1) Fredrickson, J. K.; Romine, M. F.; Beliaev, A. S.; Auchtung, J. M.;
Driscoll, M. E.; Gardner, T. S.; Nealson, K. H.; Osterman, A. L.;
Pinchuk, G.; Reed, J. L.; et al. Towards environmental systems
biology of Shewanella. Nat. Rev. Microbiol. 2008, 6 (8), 592–603.
(2) Nealson, K. H.; Saffarini, D. Iron and manganese in anaerobic
respiration: environmental significance, physiology, and regulation. Annu. Rev. Microbiol. 1994, 48, 311–43.
(3) Lovley, D. R.; Coates, J. D. Bioremediation of metal contamination. Curr. Opin. Biotechnol. 1997, 8 (3), 285–9.
(4) Logan, B. E. Exoelectrogenic bacteria that power microbial fuel
cells. Nat. Rev. Microbiol. 2009, 7 (5), 375–81.
(5) Logan, B. E.; Hamelers, B.; Rozendal, R.; Schroder, U.; Keller,
J.; Freguia, S.; Aelterman, P.; Verstraete, W.; Rabaey, K. Microbial
fuel cells: Methodology and technology. Environ. Sci. Technol.
2006, 40 (17), 5181–92.
(6) Bretschger, O.; Obraztsova, A.; Sturm, C. A.; Chang, I. S.; Gorby,
Y. A.; Reed, S. B.; Culley, D. E.; Reardon, C. L.; Barua, S.; Romine,
M. F.; et al. Current production and metal oxide reduction by
Shewanella oneidensis MR-1 wild type and mutants. Appl.
Environ. Microbiol. 2007, 73 (21), 7003–12.
(7) Marsili, E.; Baron, D. B.; Shikhare, I. D.; Coursolle, D.; Gralnick,
J. A.; Bond, D. R. Shewanella secretes flavins that mediate
extracellular electron transfer. Proc. Natl. Acad. Sci. U. S. A.
2008, 105 (10), 3968–73.
(8) Lies, D. P.; Hernandez, M. E.; Kappler, A.; Mielke, R. E.; Gralnick,
J. A.; Newman, D. K. Shewanella oneidensis MR-1 uses overlapping pathways for iron reduction at a distance and by direct
contact under conditions relevant for Biofilms. Appl. Environ.
Microbiol. 2005, 71 (8), 4414–26.
(9) Reguera, G.; McCarthy, K. D.; Mehta, T.; Nicoll, J. S.; Tuominen,
M. T.; Lovley, D. R. Extracellular electron transfer via microbial
nanowires. Nature 2005, 435 (7045), 1098–101.
(10) Gorby, Y. A.; Yanina, S.; McLean, J. S.; Rosso, K. M.; Moyles, D.;
Dohnalkova, A.; Beveridge, T. J.; Chang, I. S.; Kim, B. H.; Kim,
K. S.; et al. Electrically conductive bacterial nanowires produced
by Shewanella oneidensis strain MR-1 and other microorganisms. Proc. Natl. Acad. Sci. U. S. A. 2006, 103 (30), 11358–63.
(11) Reguera, G.; Nevin, K. P.; Nicoll, J. S.; Covalla, S. F.; Woodard,
T. L.; Lovley, D. R. Biofilm and nanowire production leads to
increased current in Geobacter sulfurreducens fuel cells. Appl.
Environ. Microbiol. 2006, 72 (11), 7345–8.
(12) Bond, D. R.; Lovley, D. R. Electricity production by Geobacter
sulfurreducens attached to electrodes. Appl. Environ. Microbiol.
2003, 69 (3), 1548–55.
(13) Srikanth, S.; Marsili, E.; Flickinger, M. C.; Bond, D. R. Electrochemical characterization of Geobacter sulfurreducens cells
immobilized on graphite paper electrodes. Biotechnol. Bioeng.
2008, 99 (5), 1065–73.
(14) Ishii, S.; Watanabe, K.; Yabuki, S.; Logan, B. E.; Sekiguchi, Y.
Comparison of electrode reduction activities of Geobacter
sulfurreducens and an enriched consortium in an air-cathode
microbial fuel cell. Appl. Environ. Microbiol. 2008, 74 (23), 7348–
55.
(15) Thormann, K. M.; Saville, R. M.; Shukla, S.; Spormann, A. M.
Induction of rapid detachment in Shewanella oneidensis MR-1
biofilms. J. Bacteriol. 2005, 187 (3), 1014–21.
(16) Torres, C. I.; Marcus, A. K.; Parameswaran, P.; Rittmann, B. E.
Kinetic experiments for evaluating the Nernst-Monod model
for anode-respiring bacteria (ARB) in a biofilm anode. Environ.
Sci. Technol. 2008, 42 (17), 6593–7.
(17) Hong, S. H.; Jeong, J.; Shim, S.; Kang, H.; Kwon, S.; Ahn, K. H.;
Yoon, J. Effect of electric currents on bacterial detachment and
inactivation. Biotechnol. Bioeng. 2008, 100 (2), 379–86.
(18) Busalmen, J. P.; de Sanchez, S. R. Changes in the electrochemical
interface as a result of the growth of Pseudomonas fluorescens
biofilms on gold. Biotechnol. Bioeng. 2003, 82 (5), 619–24.
(19) Franks, A. E.; Nevin, K. P.; Jia, H.; Izallalen, M.; Woodard, T. L.;
Lovley, D. R. Novel strategy for three-dimensional real-time
imaging of microbial fuel cell communities: monitoring the
inhibitory effects of proton accumulation within the anode
biofilm. Energy Environ. Sci. 2009, 2, 113–119.
(20) HaoYu, E.; Cheng, S.; Scott, K.; Logan, B. Microbial fuel cell
performance with non-Pt cathode catalysts. J. Power Sources
2007, 171 (2), 275–281.
(21) Dewan, A.; Beyenal, H.; Lewandowski, Z. Scaling up microbial
fuel cells. Environ. Sci. Technol. 2008, 42 (20), 7643–8.
(22) McLean, J. S.; Majors, P. D.; Reardon, C. L.; Bilskis, C. L.; Reed,
S. B.; Romine, M. F.; Fredrickson, J. K. Investigations of structure
and metabolism within Shewanella oneidensis MR-1 biofilms.
J. Microbiol. Methods 2008, 74 (1), 47–56.
(23) Beyenal, H.; Donovan, C.; Lewandowski, Z.; Harkin, G. Threedimensional biofilm structure quantification. J. Microbiol.
Methods 2004, 59 (3), 395–413.
(24) Thormann, K. M.; Saville, R. M.; Shukla, S.; Pelletier, D. A.;
Spormann, A. M. Initial phases of biofilm formation in Shewanella oneidensis MR-1. J. Bacteriol. 2004, 186 (23), 8096–
8104.
(25) Biffinger, J. C.; Pietron, J.; Ray, R.; Little, B.; Ringeisen, B. R. A
biofilm enhanced miniature microbial fuel cell using Shewanella
oneidensis DSP10 and oxygen reduction cathodes. Biosens.
Bioelectron. 2007, 22 (8), 1672–9.
(26) Ringeisen, B. R.; Henderson, E.; Wu, P. K.; Pietron, J.; Ray, R.;
Little, B.; Biffinger, J. C.; Jones-Meehan, J. M. High power density
from a miniature microbial fuel cell using Shewanella oneidensis
DSP10. Environ. Sci. Technol. 2006, 40 (8), 2629–34.
(27) McLean, J. S.; Ona, O. N.; Majors, P. D. Correlated biofilm
imaging, transport and metabolism measurements via combined nuclear magnetic resonance and confocal microscopy.
ISME J. 2008, 2 (2), 121–31.
(28) Majors, P. D.; McLean, J. S.; Fredrickson, J. K.; Wind, R. A. HMR
methods for in-situ biofilm metabolism studies: Spatial and
temporal resolved measurements. Water Sci. Technol. 2005, 52
(7), 7–12.
(29) Shaner, N. C.; Steinbach, P. A.; Tsien, R. Y. A guide to choosing
fluorescent proteins. Nat. Methods 2005, 2 (12), 905–909.
(30) Teal, T. K.; Lies, D. P.; Wold, B. J.; Newman, D. K. Spatiometabolic
stratification of Shewanella oneidensis biofilms. Appl. Environ.
Microbiol. 2006, 72 (11), 7324–30.
ES903043P
VOL. 44, NO. 7, 2010 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
9
2727