- Email: [email protected]
Miniaturizing microbial fuel cells
Review
Special Issue – Applied Microbiology
Miniaturizing microbial fuel cells Fang Qian1 and Daniel E. Morse2 1 2
Department of Chemistry and Biochemistry, University of California, Santa Cruz, CA, 95064, USA Department of Molecular, Cellular and Developmental Biology, University of California, Santa Barbara, CA, 93106-9610, USA
Microbial fuel cells (MFCs) represent an emerging technology for electricity generation from renewable biomass. Given the demand for a better understanding of the bio/inorganic interface that plays a key role in MFC energy production, small-scale MFCs are receiving considerable attention owing to their intrinsic advantages in both fundamental studies and applications as highthroughput platforms. Here, we present a brief review centered on the development of miniature MFCs at the milliliter to microliter scale. The principles, design motifs and experimental demonstrations of representative miniature MFC devices and systems are introduced, followed by a discussion of the key challenges and opportunities for realizing the exciting potentials of miniaturized MFCs. Microbial fuel cell principles Microbial fuel cells (MFCs) comprise a family of electrochemical devices that can convert biodegradable organic matter into electricity via microbial catalysis [1–5]. Compared to other fuel cell systems, the MFC employs live microorganisms to efficiently catalyze degradation of a broad range of organic substrates (most of which are abundant, non-toxic and relatively inexpensive) under mild conditions [4]. By utilizing of pathways that Nature uses to recycle the energy from renewable biomass, MFCs are operated in an environmentally benign manner, offering clean and sustainable energy. Central to MFC technology is the use of electrogenic bacterial strains that can transfer electrons produced via metabolism across the cell membrane to an external electrode [2,5,6]. This process, called extracellular electron transfer, plays a key role in harvesting bioenergy in the MFC reactor. Although the mechanism of extracellular electron transfer has not yet been fully elucidated, several possible pathways have been proposed, including direct outer membrane c-type cytochrome/anode coupling [7–9], through either redox electron mediators [10,11] or electrically conductive pili [12–14] (Figure 1). A typical MFC reactor consists of an anode and a cathode compartment that are separated by a proton exchange membrane (PEM) (Figure 1). In a working MFC, electrogenic bacterial cells are inoculated into the anode compartment, where they oxidize organic substrate present in the anolyte, generate electrons via their central metabolism, and release some of these electrons extracellularly to the anode. The extracted electrons are further Corresponding author: Morse, D.E. ([email protected]).
62
delivered through an external load to the cathode to reduce an oxidant, thus producing a current. To maintain electrochemical neutrality, protons in the anolyte will diffuse through the PEM to the cathode chamber concurrently with electron transfer. A sustainable current generation is the most basic indicator of energy conversion. The key parameters to evaluate MFC performance include current density, maximal output power density, sustainability, Coulombic efficiency, and biological oxygen demand removal efficiency. More quantitative investigation of electrode dynamics can be conducted using a potentiostat or electrochemical station [15–19]. Exploration of various device configurations [20–24], electrode materials [8,9,25–29], bacterial strains [30–33], and substrates [7,34–37] for improved MFC output are the major foci of current MFC research. The past decade has witnessed great advances in MFC technology, with ever-increasing power densities, reliabilities and diversified functionalities due largely to the evolutionary reduction in internal resistance that has been achieved by rational design of the MFC reactors. However, the output power that is produced by MFCs is still insufficient to drive most electronic devices used in our daily life. This limitation has motivated efforts in MFC research with the goal of achieving high power densities at low cost; if successful, MFCs could become a significant niche component of a balanced and sustainable energy portfolio. To achieve this goal, substantial efforts have been made in the optimization of MFC reactors with low internal resistance [22,24]; integrated or scaled-up MFC systems that can provide greater power [38]; use of MFCs for wastewater treatment [25,35,39]; exploration of new microbial strains [32,33]; and recycling diverse forms of biomass [34,36,37]. Additionally, multifunctional ‘‘MXC’’ devices have been explored extensively since 2006. These are modified MFC reactors that use the harvested bioelectricity in situ to drive other reactions, such as hydrogen production in a microbial electrolysis cell (MEC) [40] and water purification in a microbial desalination cell (MDC) [41]. Why go small? If one considers MFC reactors in terms of size, most research to date has been carried out in reactors with chamber volumes that range from several milliliters to liters and are operated on the laboratory scale. Both easy to fabricate and versatile in design, the milliliter-scale devices serve as convenient tools for exploring the interplay between device architecture and active microbes, as well as fundamental problems in electron transfer at the microbial/anode interface. Indeed, owing to the flexibility
0167-7799/$ – see front matter ß 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.tibtech.2010.10.003 Trends in Biotechnology, February 2011, Vol.29, No. 2
()TD$FIG][ Review
Trends in Biotechnology February 2011, Vol.29, No. 2
eExternal resistor Path 1 OMC
Organic matter Central metabolism
Path 2
CO2
PEM
Fe(II)
e
Pili Bacteria Path 3
Anode
Electron
H+
Fe(III) Cathode
Growth medium
mediator
Buffer TRENDS in Biotechnology
Figure 1. Working principles of a dual-chamber, ferricyanide-driven MFC. In the anode chamber, electrogenic bacterial cells catalyze the oxidation of organic matter and produce electrons (represented by black circles) in their central metabolism. Some of these electrons are transferred extracellularly to an anode via distinct pathways, including through direct outer membrane protein/anode coupling (path 1), conductive pili (path 2), and/or via self-secreted electron shuttles (path 3). Electrons are subsequently delivered through an external circuit to the cathode and used to reduce [Fe(CN)6]3 into [Fe(CN)6]4 . In accordance with each electron transfer, a proton diffuses from the anode to the cathode chamber through the proton exchange membrane. Abbreviation: OMC, outer-membrane c-type cytochrome (a group of hemecontaining membrane proteins involved in electron transfer).
in adopting favorable configurations for reducing internal resistance and improving mass transport, the best results to date have been achieved from milliliter-scale MFCs (Table 1). A 1.2-ml mini-MFC that features high surface area to volume (SAV) ratio chambers for enhanced proton diffusion has been reported, which produces a high power density of 500 W/m3 [22]. A 2.5-ml, air-breathing MFC that can generate a power density of 1010 W/m3 has also been demonstrated [24]. The appreciable power output and small volume of milliliter to liter MFCs suggest their potential use as portable power supplies, especially if connected in series to achieve increased voltage and power [38]. Microscale MFCs: entering a new regime There is a new and growing interest in the development of even smaller MFCs, in which the effective chamber volumes are reduced to the (sub)microliter regime. This
desire is motivated by the fact that we still do not have an in-depth understanding of the extracellular electron transfer processes associated with MFC operation, especially for a small colony of bacterial cells. Micro-MFCs can enable crucial studies of these processes in a smaller group of cells with excellent control over the microenvironment, thus serving as a versatile platform for fundamental MFC studies. The advantages of micro-MFCs originate from their unique structural features and scales. Prepared by microfabrication techniques (Figure 2), micro-MFCs typically feature chambers with well-defined thicknesses (tens to hundreds of micrometers) and high SAV ratios. As an immediate result, the ratio of cells coupled to the solid electrodes relative to the cells that remain in suspension increases proportionally with the SAV ratio (assuming no specific affinities or preferential partitioning), thus enrich-
Table 1. Summary of representative miniature MFCs Pmax
Imax
Anode chamber volume (ml)
Inoculum
Anode material
Projected area (cm2)
Substrate
Catholyte
Open circuit voltage (V)
2500
Mixed bacterial culture Shewanella oneidensis DSP-10 S. oneidensis MR-1 Shewanella sp. Hac353 S. oneidensis MR-1 Saccharomyces cerevisiae S. cerevisiae Shewanella putrefaciens S. oneidensis MR-1
Carbon cloth Graphite felt Gold
7
Acetate
Air
N/A
(W/m3) 1010
(mW/m2) 1800
(A/m3) 5050
(mA/m2) 9000
[24]
2
Lactate
Ferricyanide
0.7
660a
4000a
1800a
11000a
[22]
0.385
TSB
Ferricyanide
0.51
0.025*
0.4
0.34*
5.5
[47]
Gold
0.385
TSB
Air
N/A
0.17*
2.69
0.38*
6
[48]
Graphite felt Gold
25 mg
Lactate
Ferricyanide
0.65
4400b
0.2b
N/A
N/A
[49]
0.51
Glucose
Ferricyanide
0.30.5
0.5c
0.023c
1000c
150c
[51]
c
c
[52] [9]
1200 650 650 400 16 15 10 1.5
c
c
Refs.
Gold Gold
1.2 0.02
Glucose Lactate
Ferricyanide N/A
0.49 N/A
32 * N/A
4.0 N/A
2416 * 0.76
302 3.8
Gold
0.15
Lactate
Ferricyanide
0.6
16.3
1.5
1300
130
[53]
a–c
Artificial electron mediator was added: a100 mM anthraquinone-2,6-disulfonate (AQDS); b5 mM AQDS; c10 mM methylene blue. *Our calculations based on reported data. Abbreviations: N/A, not applicable; TSB, tryptic soy broth.
63
()TD$FIG][ Review
Trends in Biotechnology February 2011, Vol.29, No. 2
(a)
Si substrate
PR coating
Photolithogrpahy
Etch
(b)
Au deposition
PR coating
Photolithogrpahy
Bake
(c)
Si mold
PDMS curing
Peel off TRENDS in Biotechnology
Figure 2. Fabrication of microfluidic channels composed of various materials. (a) To create microfluidic channels in a silicon wafer, photoresist (PR) is first spin-coated on the wafer surface, followed by exposure to UV light through a photomask that specifies the microfluidic channel pattern, and selective removal of the exposed PR. The PR layer then serves as an etching mask to transfer the pattern into the silicon wafer. (b) Some polymeric PRs, such as SU-8, can be used directly to form the walls of the microfluidic chambers. First, the micro-pattern is created in the SU-8 resist (pink layer) by photolithography. The wafer is then baked to enable complete crosslinking of SU8, which turns it into a hard solid with superior chemical stability. (c) PDMS can be used to construct fine structures via soft lithography [45]. In this approach, a hard mold with surface micro-patterns serves as a template for PDMS replication. Liquid-phase PDMS pre-polymer is poured on the mold surface, which turns into a rubber-like transparent solid that can be peeled off from the mold after thermal curing.
ing the proportion of electrode-coupled cells (i.e. the cells of interest) in the microliter chambers. Conversely, because the current generation in an MFC is associated with local electron transfer at the microbial/electrode interface, the number of electrically addressable cells should scale down with electrode area. Thus, measurements can be conducted on a small collection of bacterial cells (versus a larger ensemble) in a uniform local environment offered by micro-MFCs, which would not be possible in traditional macroscopic devices. In summary, micro-MFCs could improve the tractability of fundamental investigations that aim to probe the behavior and physiology of electrogenic microorganisms and their interaction with MFCs at a new level of detail and efficiency. Integrating microfluidics with MFC research Microfluidic systems have demonstrated their utility as bioreactors in studies of a broad range of biological reactions, from fundamental biomolecular interactions and cell growth, motility and differentiation, to the production of high-purity bioproducts. Microfluidic devices (Box 1) are potentially powerful tools for investigation of microbial systems, primarily because their dimensions closely match the intrinsic scale of microbial cells. Although there have been many reports on microfluidic investigations of microorganisms [46], only a few have thus far described inves64
tigations of micro-MFCs. This is probably because the typical MFC reactor requires a relatively complicated dual-chamber configuration that is separated by an appropriate ion exchange membrane, and contains anode, cathode and well-separated anolyte and catholyte. In addition, the bulky carbon-based electrodes in conventional MFCs are not compatible with the microfabrication processes described above. Nevertheless, several groups have designed intricate microscale chambers for performing MFC research at this new dimension (Table 1). A 10-ml single-chamber reactor has been constructed using a Gortex gasket to study the electrical coupling between Shewanella putrefaciens and a surface-modified gold anode [9]. The authors have observed an extremely low background current of 0.1 nA and an instant start-up (i.e. the time interval between bacterial inoculation and the onset of current generation), which has enabled them to distinguish fine-scale changes in current generation as a function of progressive anode surface modification. In addition, milliliter-MFC arrays that are individually addressable are emerging as a versatile platform for highthroughput identification and characterization of electrochemically active microbes. Compared with conventional H-shaped, two-chamber MFCs, milliliter-MFC arrays consume less materials and reagents, enable parallel, repro-
Review Box 1. Microfluidics for manipulating the bacterial microenvironment Incubation of bacterial cells generally involves the control of a series of physical, chemical and biochemical conditions, including temperature, osmotic pressure, pH, electrochemical potential, illumination, nutrient/gas supply, removal of metabolic waste, and exclusion of contamination [42]. Conventional flask or agar-plate-based culture approaches can provide appropriate conditions for growth of large populations of cells, yet use of these methods can be difficult (and sometimes impossible) when precision of control in the microenvironment around individual cells is required. In this regard, microfluidic technologies, which offer a powerful tool to manipulate fluids in a predicable manner at micrometer dimensions [43], can provide unprecedented control of the cellular microenvironment. Microfluidic systems are typically composed of well-defined, micro- to nanoliter channels or chambers on a surface. These are not simply scaled-down versions of macrofluidic devices, because fluid behavior changes significantly when the channel/ chamber size is reduced to a critical value (tens to hundreds of microns depending on specific conditions) [44,45]. Uniform chemical conditions are maintained in each flow layer, while mass transport between the layers is diffusion-limited. Fluid dynamics in microfluidic systems can be predetermined by patterned channels and specially designed micro-components (e.g. valves and pumps), and therefore, is highly predictable and controllable. Microfluidic systems can be constructed by well-established microfabrication processes, as exemplified by photolithography. In brief, photolithography is used to transfer a computer-designed pattern from a mask to a substrate mediated by photosensitive polymer (PR). The topographically patterned substrate can subsequently be used as templates for preparing a variety of microfluidic systems. Figure 2 briefly outlines the technical procedures in fabrication of three representative types of microfluidic chambers, made of silicon, PR and PDMS, respectively.
ducible and cost-effective analysis of electrogenic strains, and thus can greatly accelerate MFC research. For example, a 24-well MFC array has been developed using gold as anode material [47,48]. Each well has an anode chamber volume of about 650 ml and functions as an independent MFC. The MFC array prototype has been used to screen environmental microbes. Using this array, researchers have succeeded in isolating a bacterial strain that displays 2.3-fold higher power output than the Shewanella oneidensis MR-1 reference strain. In another case, multi-anode/ common cathode MFC arrays have been created by employing commercially available 1-ml pipette tips as MFC chambers, each with an anode chamber volume of 400 ml [49]. The resultant array of micro-MFCs allows for efficient monitoring of bacterial growth cycles and comparison of carbon source utilization of S. oneidensis MR-1 species. Besides the micro-chambered MFCs with fixed-volume fluids, micro-MFCs that contain microfluidic channels that allow fluid exchange have also been developed. The earliest report was a silicon-based microfluidic MFC system [50]. Two silicon wafers were first micro-machined to embed convoluted microfluidic channels, followed by wet-etch to form through-holes that served as fluidic inlets and outlets. Using an evaporated gold layer as electrodes, two wafers were pressed face-to-face, yet were separated by a PEM film. Glucose was used as the carbon source. Relative to the negative control, inoculation of Saccharomyces cerevisiae in this 16-ml MFC yielded an apparent current of 15 mA/ cm2 (on a 10-V load for 14 min), which corresponded to a
Trends in Biotechnology February 2011, Vol.29, No. 2
power output of 0.5 W/m3. Using a modified micro-MFC topped with transparent glass covers, the authors later reported current generation from a photosynthetic strain of Cyanophyta [51]. The new micro-MFC generated a current via photosynthesis when illuminated, and via glucose fermentation in the dark. Under a 10-V load, the power density was 40 pW/cm2. In a more recent study, a 15-ml polydimethylsiloxane (PDMS)-based micro-MFC with micropillar anodes has been demonstrated [52]. By using the same strain (S. cerevisiae) and a 1-kV load, the optimized device produced an increased current of 4.3 mA/ cm2, and a power of 42.4 nW/cm2 (3.4 W/m3, as calculated from the reported data), with output sustained for 60 min. Micro-MFC case study: Shewanella Here, we take our recently developed 1.5-ml MFC device as an example to discuss the fabrication and measurement of micro-MFCs [53]. This dual-chamber microfluidic MFC allows on-chip bacterial culture and conversion of bacterial metabolism into electricity, by utilization of the smallest anode chamber for all reported MFCs. As shown in Figure 3a and b, the micro-MFC was constructed by sequentially stacking a silicon wafer prefabricated with a gold-film anode and an SU-8 microfluidic channel (1.5 ml), a PEM, a carbon-cloth cathode, and a PDMS cathode chamber (4 ml). The device assembly was sandwiched between two acrylic plates. The well-defined channel thicknesses (100 mm) provided a short distance between the electrodes, thereby allowing efficient proton diffusion while reducing the electrolyte resistance. Inoculation of S. oneidensis MR-1 in the MFC anode chamber yielded a pronounced current density of 1.3 mA/cm2 on a 100-V resistor, with current sustained for 20 h. The current generation was reproducible when operated in a batchfed mode (Figure 3c). The Coulombic efficiency of the MFC that consumed lactate was calculated to be 2.8% – much lower than macroscopic MFCs (up to 90% [54]) – which indicates that the chemical-to-electricity conversion has not yet been optimized and represents an area that requires improvement. Electron microscopy images have shown that the MR-1 cells grow on the gold anode with high density and in a multi-layered arrangement (Figure 3c, inset). Remarkably, comparative studies of bacterial growth in various reactors have revealed an enhanced cell density on the gold anode in a 1.5-ml micro-MFC (>2 106 cell/mm2) compared to that in a 5-ml Petri dish (5 104 cell/mm2). These results were consistent with the increased current density and short start-up phase of micro-MFCs [9,50–53], which suggests that cell-to-anode coupling was enhanced in the micro-chamber environments. Polarization and power curves have revealed that the micro-MFC (Figure 3d) operates in a region where Ohmic loss was dominant [55] and the maximal power generated was 23 nW (15.3 W/m3). Linear fitting of the polarization curve yielded a relatively high internal resistance of 13 kV, which is orders of magnitude higher than that of the optimized milliliter-MFCs (down to several V) [23,56]. The high internal resistance could have stemmed from the small electrode area, polarization loss owing to slow electrode reactions, and/or electrolyte/PEM resistances 65
()TD$FIG][ Review
Trends in Biotechnology February 2011, Vol.29, No. 2
(a)
K3Fe(CN)6
(b) Catholyte 1
PDMS
2
Carbon cloth PEM
Anolyte
Growth medium
R
+ SU-8
3
C MF
Gold
ro-
Mic
(c)
Si wafer
2 cm
(d)
2.0
50
25
40
20
30
15
20
10
10
5
0
0
Voltage (mV)
1.0
Power (nW)
Current (µA)
1.5
Lactate
0.5
0.0 0
20
40
60
80
100 120 140 160 180
Time (h)
0
0.5
1.0
1.5
2.0
Current (µA) TRENDS in Biotechnology
Figure 3. Fabrication and performance of a 1.5-ml MFC device. (a) Schematic presentation of the MFC design and the key components. (b) Photograph of the microfluidic MFC device described in (a), filled with electrolytes. (c) Current recorded from a batch-fed micro-MFC over time. The micro-MFC generated current in response to 20 mM lactate addition (indicated by arrows) and sustained the current for 20–30 h. Inset: scanning electron microscopy image of the bacteria colony at the anode. Scale bar, 2 mm. (d) Polarization (solid symbols) and power (empty symbols) curves measured from the micro-MFC device. Reproduced with permission from Ref. [53].
that scaled inversely with the electrode area. The observed high internal resistance limits power output and represents a major challenge for all (sub)microliter-scale MFCs [56]. To solve this problem, the development of new electrode materials [25,27,57] with high surface area, low resistance, and strong affinity for bacterial cells (facilitating extracellular electron transfer) is crucial. To date, the potential of utilizing microfluidic devices for MFC research has not been fully explored, although such devices have demonstrated several general advantages over conventional MFC devices. As a result of the size reduction, micro-MFCs show low background signals, rapid start-up, and high sensitivity to bacterial metabolism. Most importantly, the much smaller electrode in the microMFC is more responsive to a local electrochemical change in its microchamber than a macroscopic device, which might allow MFC research to enter the realm of measurement of electrogenesis by a single bacterial cell. Key challenges for miniaturized MFCs The development of micro-MFCs offers new and unique opportunities to analyze MFC reactions at small scales in a well-controlled microenvironment. However, miniaturization of the electrodes and the reaction chamber also present 66
challenges for micro-MFCs. One of the major challenges is to reduce their large internal resistance, which, if it can be solved, could lead to better output power densities and potential applications as power supplies for ultra-small electronics. Theoretically, the membrane and electrolyte resistances can be reduced by carefully designed reactor architectures; for example, by designing chambers with even higher SAV ratios, using a thinner PEM, and increasing the ionic concentration of electrolytes to the limits tolerated by the bacterial cells [55]. Systematic exploration of growth conditions and alternative strains could help to identify the best bacterial candidates for proliferation and electrogenesis in the microfluidic environment that can differ significantly from that of conventional cultivation conditions. Towards reducing resistance associated with the electrodes, incorporation of high-surface-area, carbon-based electrodes in the microscale chambers might offer a promising solution. Our recent result of a 4-ml carbon clothanode MFC exhibited enhanced power output (250 nW) and power density (62.5 W/m3) compared to the 1.5-ml goldanode MFC (23 nW, 15 W/m3) operated under identical conditions (D. Morse, unpublished). It is desirable to explore aggressively non-conventional nanoscale electrodes, such as carbon nanotubes, graphene or semiconductor
Review nanowires, for use in micro-MFCs. Additionally, one also must pay attention to the design of the cathode half-cells to prevent cathode limitation. Another key scientific challenge is the coupling of fundamental electrochemical studies to the microfluidic MFC platform. Although rational design of micro-reactors can lead to an optimized solution for a micro-MFC with good power densities, analytical electrochemical measurements provide complementary information for a quantitative understanding of the electrode kinetics, bacterial redox potential, mass transport, and internal resistance distribution. However, most important electrochemical measurements presently require the use of a bulky reference electrode, which cannot yet be coupled into a microscale chamber. Despite the fact that planar Ag/AgCl microelectrodes have been demonstrated as reference electrodes elsewhere, their stabilities and lifetimes are still insufficient to support long-term MFC studies [58]. Therefore, the development of a reliable three-electrode microelectrochemical system would be crucial for enabling a series of fundamental electrochemical characterizations in microfluidic MFCs. Future prospects Compared to the well-established milli- and macro-MFCs, micro-MFCs are still in their infancy. Although the small power output and high fabrication cost exclude microMFCs from large-scale use in power supply and waterpurification applications, their unique dimensions provide advantages for microfluidic MFCs in several novel areas. Ultra compact and fast current-based assays High-density integration of functional micro-MFCs could allow researchers to conduct multiple electrical, electrochemical or biochemical measurements in parallel. Considering a micro-MFC array composed of MFC units with a side-length of 100 mm and a spacing of 100 mm, 2500 individually operated MFCs can be arrayed within a 1cm2 area. As discussed above, high-SAV-ratio microscale reactors can shorten the ion diffusion length and promote biofilm development to convert analytes (substrates) more efficiently into detectable current signals. Such arrays of microliter-volume MFCs thus offer unique advantages of fast start-up, high sensitivity and superior microfluidic control over the measured microenvironment, which makes them good candidates for rapid screening of electrode materials, bacterial strains and growth media. Thus, for instance, the micro-MFC array can be used to select optimally efficient electrogenic bacterial strains, which can then be used in conventional MFC reactors on larger scales. Bioenergy sources for ultra-small electronics With the development of nanoscience and technology, a broad range of nanodevices have been demonstrated for the next-generation applications in computing, communication and sensing [59,60]. These nanoscale electronic elements and networks generate, process and transmit signals on a low-energy basis (nW to mW) [59], which can in principle be supplied by micro-MFCs. Through on-chip integration, the combination of microbial energy
Trends in Biotechnology February 2011, Vol.29, No. 2
supply and nanodevice operation could lead to self-sustained nanosystems that might find significant uses in diverse areas. Further studies are still needed to test whether the long-term operation of micro-MFCs is practical, and how to overcome potential challenges, such as cells clogging the microchannel. Real-time visualization of electrogenic cells in an MFC The microscale thickness of micro-MFCs is compatible with the working distances of conventional optical microscopes and thus can be directly placed under an optical microscope for observing live bacterial behavior. The ability to visualize live cells in situ in an MFC requires device transparency for light transmission, which can be easily obtained by PDMS- or glass-based micro-MFCs. In addition, an enhanced optical contrast between cells and the environment is necessary for live-cell investigation, which can be achieved using staining dyes [61], genetically modified fluorescent strains [61,62] or phase-contrast imaging [63]; albeit, these requirements might add to the complexity of the reactor. Correlation of optical and electrical/ electrochemical studies of MFCs could provide complementary information on cell growth and functioning in a microMFC, and this in turn could suggest novel strategies for unravelling and optimizing the details of extracellular electron transfer mechanism in MFCs. Single-bacterium studies One of the most attractive promises of micro-MFCs is to study a single bacterium, which could revolutionize our understanding in these electricity-generating microorganisms. To date, the MFC field has largely focused on the collaborative performance of a large population of bacterial cells; typically in the form of a biofilm. The biofilm is a complex mixture of dead, live and dividing cells, embedded within a self-produced matrix of extracellular polymeric substances [64,65]. Mass transport and electron transfer in the biofilm is unknown, complex and uncontrolled. Measurements of the electrogenic performance of biofilms yield averaged results from diverse and changing populations, which makes it difficult to achieve reproducibility and to obtain precise, quantitative information on specific events in the microbial electron transfer chain. By contrast, single-bacterium analyses could allow us to elucidate this basic information from individual bacterial cells in a controlled microenvironment, which could provide a higher-resolution perspective in understanding the key electrochemical and biochemical processes at the bacterial/ electrode interface. Only with further in-depth comprehension of these fundamental processes will we be able to improve the efficiency of bacterial electrogenesis and further advance MFC research and development. Figure 4 illustrates a possible design of a single-bacterium microfluidic MFC device. Microelectrodes are first evaporated onto the substrate and coupled in the microfluidic channel. These microelectrodes are individually addressable, serve as anodes, and share a common PEC/ cathode assembly, thus forming a microelectrode MFC array. On this unique platform, the maximal number of bacterial cells coupled to the anode can be controlled by individual anode size. Intercellular communication among 67
()TD$FIG][ Review
Trends in Biotechnology February 2011, Vol.29, No. 2
Microfluidic channel
Fluid out
Fluid in
Microelectrode
TRENDS in Biotechnology
Figure 4. Artistic rendering of a single-bacterium micro-MFC device. Microelectrode arrays are defined within the microfluidic anode chamber. By control of the electrode size, spacing and cell density, such a device should make it possible to probe electricity generation from a single electrogenic cell.
bacterial cells (e.g. quorum sensing) could be studied as a function of cell density with such a device. The ability to create complex electrode patterns with well-defined morphologies (e.g. pillars, trenches and holes), combined with the precise control of electrode position, could be used to tune the 3D geometry of the microbial/anode interface at the (sub)microscale. Moreover, the incorporation of nanoscale electrodes, such as nanowire arrays, in the microfluidic platform might enable multi-point, intracellular measurements of a single bacterium, which introduces further exciting opportunities in fundamental MFC research. Acknowledgements We thank Dr. Ning Cao at the University of California at Santa Barbara Nanofabrication Center for valuable discussions. This work was supported in part by grants from the U.S. Department of Energy (grant # DE-FG02-02ER46006) and the U.S. Army Research Office (through contract no. W911NF-09-D-0001 to the Institute for Collaborative Biotechnologies). F. Q. thanks the support in part of this work by the National Science Foundation (award # CBET 1034222).
References 1 Bond, D.R. et al. (2002) Electrode-reducing microorganisms harvesting energy from marine sediments. Science 295, 483–495 2 Lovley, D.R. (2006) Bug juice: harvesting electricity with microorganisms. Nat. Rev. Microbiol. 4, 497–508 3 Logan, B.E. et al. (2006) Microbial fuel cells: methodology and technology. Environ. Sci. Technol. 40, 5181–5192 4 Logan, B.E. (2008) Microbial Fuel Cells, John Wiley 5 Fredrickson, J.K. et al. (2008) Towards environmental systems biology of Shewanella. Nat. Rev. Microbiol. 6, 592–603 6 Bretschger, O. et al. (2007) Current production and metal oxide reduction by Shewanella oneidensis MR-1 wild type and mutants. Appl. Environ. Microbiol. 73, 7003–7012 7 Chaudhuri, S.K. and Lovley, D.R. (2003) Electricity generation by direct oxidation of glucose in mediatorless microbial fuel cells. Nat. Biotechnol. 21, 1229–1232 8 Bond, D.R. and Lovley, D.R. (2003) Electricity production by Geobacter sulfurreducens attached to gold electrodes. Appl. Environ. Microbiol. 69, 1548–1555 9 Crittenden, S.R. et al. (2006) Mediating electron transfer from bacteria to a gold electrode via a self-assembled monolayer. Langmuir 22, 9473–9476 10 Newman, D.K. and Kolter, R. (2000) A role for excreted quinones in extracellular electron transfer. Nature 405, 94–97
68
11 Marsili, E. et al. (2008) Shewanella secretes flavins that mediate extracellular electron transfer. Proc. Natl. Acad. Sci. U. S. A. 105, 3968–3973 12 Reguera, G. et al. (2005) Extracellular electron transfer via microbial nanowires. Nature 435, 1098–1101 13 Gorby, Y.A. et al. (2006) Electrically conductive bacterial nanowires produced by Shewanella oneidensis strain MR-1 and other microorganisms. Proc. Natl. Acad. Sci. U. S. A. 103, 11358–11363 14 Reguera, G. et al. (2006) Biofilm and nanowire production leads to increased current in Geobacter sulfurreducens fuel cells. Appl. Environ. Microbiol. 72, 7345–7348 15 Pessanha, M. et al. (2004) Redox characterization of Geobacter sulfurreducens cytochrome c7: physiological relevance of the conserved residue F15 probed by site-specific mutagenesis. Biochemistry 43, 9909–9917 16 Rabaey, K. et al. (2007) Microbial ecology meets electrochemistry: electricity-driven and driving communities. Int. Soc. Microb. Ecol. J. 1, 9–18 17 Aelterman, P. et al. (2008) The anode potential regulates bacterial activity in microbial fuel cells. Appl. Microbiol. Biotechnol. 78, 409–418 18 Firer-Sherwood, M. et al. (2008) Electrochemical interrogations of the Mtr cytochromes from Shewanella: opening a potential window. J. Biol. Inorg. Chem. 3, 849–854 19 He, Z. and Mansfeld, F. (2009) Exploring the use of electrochemical impedance spectroscopy in microbial fuel cell studies. Energy Environ. Sci. 2, 215–219 20 Liu, H. et al. (2004) Production of electricity during wastewater treatment using a single chamber microbial fuel cell. Environ. Sci. Technol. 38, 2281–2285 21 Cheng, S. et al. (2006) Increased power generation in a continuous flow MFC with advective flow through the porous anode and reduced electrode spacing. Environ. Sci. Technol. 40, 2426–2432 22 Ringeisen, B.R. et al. (2006) High power density from a miniature microbial fuel cell using Shewanella oneidensis DSP10. Environ. Sci. Technol. 40, 2629–2634 23 He, Z. et al. (2006) The upflow microbial fuel cell with an interior cathode: assessment of the internal resistance by impedance spectroscopy. Environ. Sci. Technol. 40, 5212–5217 24 Fan, Y.Z. et al. (2007) Enhanced Coulombic efficiency and power density of air-cathode microbial fuel cells with an improved cell configuration. J. Power Sources 171, 348–354 25 Dumitru, A. et al. (2008) Biofilm growth from wastewater on MWNTs and carbon aerogels. Physica. Status Solidi (a) 205, 1484–1487 26 Richter, H. et al. (2008) Electricity generation by Geobacter sulfurreducens attached to gold electrodes. Langmuir 24, 4376–4379 27 Qiao, Y. et al. (2008) Nanostructured polyanifine/titanium dioxide composite anode for microbial fuel cells. ACS Nano. 2, 113–119 28 Wang, X. et al. (2009) Use of carbon mesh anodes and the effect of different pretreatment methods on power production in microbial fuel cells. Environ. Sci. Technol. 43, 6870–6874 29 Liu, Y. et al. (2010) The study of electrochemically active biofilms on different carbon-based materials in microbial fuel cells. Biosens. Bioelectron. 25, 2167–2171 30 Rosenbaum, M. et al. (2010) Light energy to bioelectricity: photosynthetic microbial fuel cells. Curr. Opin. Biotechnol. 21, 259–264 31 Izallalen, M. et al. (2008) Geobacter sulfurreducens strain engineered for increased rates of respiration. Metab. Eng. 10, 267–275 32 Xing, D.F. et al. (2008) Electricity generation by Rhodopseudomonas palustris DX-1. Environ. Sci. Technol. 42, 4146–4151 33 Yi, H. et al. (2009) Selection of a variant of Geobacter sulfurreducens with enhanced capacity for current production in microbial fuel cells. Biosens. Bioelectron. 24, 3498–3503 34 Gil, G.C. et al. (2003) Energy from algae using microbial fuel cells. Biotechnol. Bioeng. 103, 1068–1076 35 Min, B. and Logan, B.E. (2004) Continuous electricity generation from domestic wastewater and organic substrates in a flat plate microbial fuel cell. Environ. Sci. Technol. 38, 5809–5814 36 Selembo, P.A. et al. (2009) High hydrogen production from glycerol or glucose by electrohydrogenesis using microbial electrolysis cells. Int. J. Hydrogen Energy 34, 5373–5381 37 Lalaurette, E. et al. (2009) Hydrogen production from cellulose in a twostage process combining fermentation and electrohydrogenesis. Int. J. Hydrogen Energy 34, 6201–6210
Review 38 Dekke, A. et al. (2009) Analysis and improvement of a scaled-up and stacked microbial fuel cell. Environ. Sci. Technol. 43, 9038–9042 39 He, Z. et al. (2005) Electricity generation from artificial wastewater using an upflow microbial fuel cell. Environ. Sci. Technol. 39, 5262– 5267 40 Cheng, S. and Logan, B.E. (2007) Sustainable and efficient biohydrogen production via electrohydrogenesis. Proc. Natl. Acad. Sci. U. S. A. 104, 18871–18873 41 Cao, X. et al. (2009) A new method for water desalination using microbial desalination cells. Environ. Sci. Technol. 43, 7148–7152 42 Wilkinson, J.F. (1972) Introduction to Microbiology, Blackwell Scientific 43 Whitesides, G.M. (2006) The origins and the future of microfluidics. Nature 442, 368–373 44 Atencia, J. and Beebe, D.J. (2005) Controlled microfluidic interfaces. Nature 437, 648–655 45 Whitesides, G.M. et al. (2001) Soft lithography in biology and biochemistry. Annu. Rev. Biomed. Eng. 3, 335–373 46 Weibel, D.B. et al. (2009) Microfabrication meets microbiology. Nat. Rev. Microbiol. 5, 209–218 47 Hou, H. et al. (2009) Microfabricated microbial fuel cell arrays reveal electrochemically active microbes. PLoS ONE 4, e6570 48 Hou, H. et al. (2011) Air-cathode microbial fuel cell array: a device for identifying and characterizing electrochemically active microbes. Biosens. Bioelectron. 26, 2680–2684 49 Biffinger, J.C. et al. (2009) Characterization of electrochemically active bacteria utilizing a high-throughput voltage-based screening assay. Biotechnol. Bioeng. 102, 436–444 50 Chiao, M. et al. (2003) Micromachined microbial fuel cells, In Proceedings of 16th IEEE MicroElectro Mechanical Systems Conference, Kyoto, Japan. pp. 383–386 51 Chiao, M. et al. (2006) Micromachined microbial and photosynthetic fuel cells. J. Micromech. Microeng. 16, 2547–2553
Trends in Biotechnology February 2011, Vol.29, No. 2 52 Siu, C. and Chiao, M. (2008) A microfabricated PDMS microbial fuel cell. J. Micro Electrochem. Syst. 17, 1329–1341 53 Qian, F. et al. (2009) A 1.5-microliter microbial fuel cell for on-chip bioelectricity generation. Lab. Chip. 9, 3076–3081 54 Chang, I.S. et al. (2004) Continuous determination of biochemical oxygen demand using microbial fuel cell type biosensor. Biosens. Bioelectron. 19, 607–613 55 Fan, Y. et al. (2008) Quantification of the internal resistance distribution of microbial fuel cells. Environ. Sci. Technol. 42, 8101–8107 56 Logan, B.E. and Regan, J.M. (2006) Microbial fuel cells – challenges and applications. Environ. Sci. Technol. 40, 5172–5180 57 Peng, L. et al. (2010) Carbon nanotubes as electrode modifier promoting direct electron transfer from Shewanella oneidensis. Biosens. Bioelectron. 25, 1248–1251 58 Polk, B.J. et al. (2006) Ag/AgCl microelectrodes with improved stability for microfluidics. Sens. Actuators B Chem. 114, 239–247 59 Li, Y. et al. (2006) Nanowire electronics and optoelectronic devices. Mater. Today 9, 18–27 60 Lieber, C.M. and Wang, Z.L. (2007) Functional nanowires. MRS Bull. 32, 99–104 61 Franks, A.E. et al. (2010) Microtoming coupled to microarray analysis to evaluate the spatial metabolic status of Geobacter sulfurreducens biofilms. Int. Soc. Microb. Ecol. J. 4, 509–519 62 Teal, T.K. et al. (2006) Spatiometabolic stratification of Shewanella oneidensis biofilms. Appl. Microbiol. Biotechnol. 72, 7324–7330 63 Jiang, X. et al. (2010) Probing electron transfer mechanisms in Shewanella oneidensis MR-1 using a nanoelectrodes platform and single-cell imaging. Proc. Natl. Acad. Sci. U. S. A. 107, 16806–16810 64 Dunne Jr, W.M. (2002) Bacterial adhesion: seen any good biofilms lately. Clin. Microbiol. Rev. 15, 155–166 65 Biffinger, J.C. et al. (2007) A biofilm enhanced miniature microbial fuel cell using Shewanella oneidensis DSP10 and oxygen reduction cathodes. Biosens. Bioelectron. 22, 1672–1679
69
Special Issue – Applied Microbiology
Miniaturizing microbial fuel cells Fang Qian1 and Daniel E. Morse2 1 2
Department of Chemistry and Biochemistry, University of California, Santa Cruz, CA, 95064, USA Department of Molecular, Cellular and Developmental Biology, University of California, Santa Barbara, CA, 93106-9610, USA
Microbial fuel cells (MFCs) represent an emerging technology for electricity generation from renewable biomass. Given the demand for a better understanding of the bio/inorganic interface that plays a key role in MFC energy production, small-scale MFCs are receiving considerable attention owing to their intrinsic advantages in both fundamental studies and applications as highthroughput platforms. Here, we present a brief review centered on the development of miniature MFCs at the milliliter to microliter scale. The principles, design motifs and experimental demonstrations of representative miniature MFC devices and systems are introduced, followed by a discussion of the key challenges and opportunities for realizing the exciting potentials of miniaturized MFCs. Microbial fuel cell principles Microbial fuel cells (MFCs) comprise a family of electrochemical devices that can convert biodegradable organic matter into electricity via microbial catalysis [1–5]. Compared to other fuel cell systems, the MFC employs live microorganisms to efficiently catalyze degradation of a broad range of organic substrates (most of which are abundant, non-toxic and relatively inexpensive) under mild conditions [4]. By utilizing of pathways that Nature uses to recycle the energy from renewable biomass, MFCs are operated in an environmentally benign manner, offering clean and sustainable energy. Central to MFC technology is the use of electrogenic bacterial strains that can transfer electrons produced via metabolism across the cell membrane to an external electrode [2,5,6]. This process, called extracellular electron transfer, plays a key role in harvesting bioenergy in the MFC reactor. Although the mechanism of extracellular electron transfer has not yet been fully elucidated, several possible pathways have been proposed, including direct outer membrane c-type cytochrome/anode coupling [7–9], through either redox electron mediators [10,11] or electrically conductive pili [12–14] (Figure 1). A typical MFC reactor consists of an anode and a cathode compartment that are separated by a proton exchange membrane (PEM) (Figure 1). In a working MFC, electrogenic bacterial cells are inoculated into the anode compartment, where they oxidize organic substrate present in the anolyte, generate electrons via their central metabolism, and release some of these electrons extracellularly to the anode. The extracted electrons are further Corresponding author: Morse, D.E. ([email protected]).
62
delivered through an external load to the cathode to reduce an oxidant, thus producing a current. To maintain electrochemical neutrality, protons in the anolyte will diffuse through the PEM to the cathode chamber concurrently with electron transfer. A sustainable current generation is the most basic indicator of energy conversion. The key parameters to evaluate MFC performance include current density, maximal output power density, sustainability, Coulombic efficiency, and biological oxygen demand removal efficiency. More quantitative investigation of electrode dynamics can be conducted using a potentiostat or electrochemical station [15–19]. Exploration of various device configurations [20–24], electrode materials [8,9,25–29], bacterial strains [30–33], and substrates [7,34–37] for improved MFC output are the major foci of current MFC research. The past decade has witnessed great advances in MFC technology, with ever-increasing power densities, reliabilities and diversified functionalities due largely to the evolutionary reduction in internal resistance that has been achieved by rational design of the MFC reactors. However, the output power that is produced by MFCs is still insufficient to drive most electronic devices used in our daily life. This limitation has motivated efforts in MFC research with the goal of achieving high power densities at low cost; if successful, MFCs could become a significant niche component of a balanced and sustainable energy portfolio. To achieve this goal, substantial efforts have been made in the optimization of MFC reactors with low internal resistance [22,24]; integrated or scaled-up MFC systems that can provide greater power [38]; use of MFCs for wastewater treatment [25,35,39]; exploration of new microbial strains [32,33]; and recycling diverse forms of biomass [34,36,37]. Additionally, multifunctional ‘‘MXC’’ devices have been explored extensively since 2006. These are modified MFC reactors that use the harvested bioelectricity in situ to drive other reactions, such as hydrogen production in a microbial electrolysis cell (MEC) [40] and water purification in a microbial desalination cell (MDC) [41]. Why go small? If one considers MFC reactors in terms of size, most research to date has been carried out in reactors with chamber volumes that range from several milliliters to liters and are operated on the laboratory scale. Both easy to fabricate and versatile in design, the milliliter-scale devices serve as convenient tools for exploring the interplay between device architecture and active microbes, as well as fundamental problems in electron transfer at the microbial/anode interface. Indeed, owing to the flexibility
0167-7799/$ – see front matter ß 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.tibtech.2010.10.003 Trends in Biotechnology, February 2011, Vol.29, No. 2
()TD$FIG][ Review
Trends in Biotechnology February 2011, Vol.29, No. 2
eExternal resistor Path 1 OMC
Organic matter Central metabolism
Path 2
CO2
PEM
Fe(II)
e
Pili Bacteria Path 3
Anode
Electron
H+
Fe(III) Cathode
Growth medium
mediator
Buffer TRENDS in Biotechnology
Figure 1. Working principles of a dual-chamber, ferricyanide-driven MFC. In the anode chamber, electrogenic bacterial cells catalyze the oxidation of organic matter and produce electrons (represented by black circles) in their central metabolism. Some of these electrons are transferred extracellularly to an anode via distinct pathways, including through direct outer membrane protein/anode coupling (path 1), conductive pili (path 2), and/or via self-secreted electron shuttles (path 3). Electrons are subsequently delivered through an external circuit to the cathode and used to reduce [Fe(CN)6]3 into [Fe(CN)6]4 . In accordance with each electron transfer, a proton diffuses from the anode to the cathode chamber through the proton exchange membrane. Abbreviation: OMC, outer-membrane c-type cytochrome (a group of hemecontaining membrane proteins involved in electron transfer).
in adopting favorable configurations for reducing internal resistance and improving mass transport, the best results to date have been achieved from milliliter-scale MFCs (Table 1). A 1.2-ml mini-MFC that features high surface area to volume (SAV) ratio chambers for enhanced proton diffusion has been reported, which produces a high power density of 500 W/m3 [22]. A 2.5-ml, air-breathing MFC that can generate a power density of 1010 W/m3 has also been demonstrated [24]. The appreciable power output and small volume of milliliter to liter MFCs suggest their potential use as portable power supplies, especially if connected in series to achieve increased voltage and power [38]. Microscale MFCs: entering a new regime There is a new and growing interest in the development of even smaller MFCs, in which the effective chamber volumes are reduced to the (sub)microliter regime. This
desire is motivated by the fact that we still do not have an in-depth understanding of the extracellular electron transfer processes associated with MFC operation, especially for a small colony of bacterial cells. Micro-MFCs can enable crucial studies of these processes in a smaller group of cells with excellent control over the microenvironment, thus serving as a versatile platform for fundamental MFC studies. The advantages of micro-MFCs originate from their unique structural features and scales. Prepared by microfabrication techniques (Figure 2), micro-MFCs typically feature chambers with well-defined thicknesses (tens to hundreds of micrometers) and high SAV ratios. As an immediate result, the ratio of cells coupled to the solid electrodes relative to the cells that remain in suspension increases proportionally with the SAV ratio (assuming no specific affinities or preferential partitioning), thus enrich-
Table 1. Summary of representative miniature MFCs Pmax
Imax
Anode chamber volume (ml)
Inoculum
Anode material
Projected area (cm2)
Substrate
Catholyte
Open circuit voltage (V)
2500
Mixed bacterial culture Shewanella oneidensis DSP-10 S. oneidensis MR-1 Shewanella sp. Hac353 S. oneidensis MR-1 Saccharomyces cerevisiae S. cerevisiae Shewanella putrefaciens S. oneidensis MR-1
Carbon cloth Graphite felt Gold
7
Acetate
Air
N/A
(W/m3) 1010
(mW/m2) 1800
(A/m3) 5050
(mA/m2) 9000
[24]
2
Lactate
Ferricyanide
0.7
660a
4000a
1800a
11000a
[22]
0.385
TSB
Ferricyanide
0.51
0.025*
0.4
0.34*
5.5
[47]
Gold
0.385
TSB
Air
N/A
0.17*
2.69
0.38*
6
[48]
Graphite felt Gold
25 mg
Lactate
Ferricyanide
0.65
4400b
0.2b
N/A
N/A
[49]
0.51
Glucose
Ferricyanide
0.30.5
0.5c
0.023c
1000c
150c
[51]
c
c
[52] [9]
1200 650 650 400 16 15 10 1.5
c
c
Refs.
Gold Gold
1.2 0.02
Glucose Lactate
Ferricyanide N/A
0.49 N/A
32 * N/A
4.0 N/A
2416 * 0.76
302 3.8
Gold
0.15
Lactate
Ferricyanide
0.6
16.3
1.5
1300
130
[53]
a–c
Artificial electron mediator was added: a100 mM anthraquinone-2,6-disulfonate (AQDS); b5 mM AQDS; c10 mM methylene blue. *Our calculations based on reported data. Abbreviations: N/A, not applicable; TSB, tryptic soy broth.
63
()TD$FIG][ Review
Trends in Biotechnology February 2011, Vol.29, No. 2
(a)
Si substrate
PR coating
Photolithogrpahy
Etch
(b)
Au deposition
PR coating
Photolithogrpahy
Bake
(c)
Si mold
PDMS curing
Peel off TRENDS in Biotechnology
Figure 2. Fabrication of microfluidic channels composed of various materials. (a) To create microfluidic channels in a silicon wafer, photoresist (PR) is first spin-coated on the wafer surface, followed by exposure to UV light through a photomask that specifies the microfluidic channel pattern, and selective removal of the exposed PR. The PR layer then serves as an etching mask to transfer the pattern into the silicon wafer. (b) Some polymeric PRs, such as SU-8, can be used directly to form the walls of the microfluidic chambers. First, the micro-pattern is created in the SU-8 resist (pink layer) by photolithography. The wafer is then baked to enable complete crosslinking of SU8, which turns it into a hard solid with superior chemical stability. (c) PDMS can be used to construct fine structures via soft lithography [45]. In this approach, a hard mold with surface micro-patterns serves as a template for PDMS replication. Liquid-phase PDMS pre-polymer is poured on the mold surface, which turns into a rubber-like transparent solid that can be peeled off from the mold after thermal curing.
ing the proportion of electrode-coupled cells (i.e. the cells of interest) in the microliter chambers. Conversely, because the current generation in an MFC is associated with local electron transfer at the microbial/electrode interface, the number of electrically addressable cells should scale down with electrode area. Thus, measurements can be conducted on a small collection of bacterial cells (versus a larger ensemble) in a uniform local environment offered by micro-MFCs, which would not be possible in traditional macroscopic devices. In summary, micro-MFCs could improve the tractability of fundamental investigations that aim to probe the behavior and physiology of electrogenic microorganisms and their interaction with MFCs at a new level of detail and efficiency. Integrating microfluidics with MFC research Microfluidic systems have demonstrated their utility as bioreactors in studies of a broad range of biological reactions, from fundamental biomolecular interactions and cell growth, motility and differentiation, to the production of high-purity bioproducts. Microfluidic devices (Box 1) are potentially powerful tools for investigation of microbial systems, primarily because their dimensions closely match the intrinsic scale of microbial cells. Although there have been many reports on microfluidic investigations of microorganisms [46], only a few have thus far described inves64
tigations of micro-MFCs. This is probably because the typical MFC reactor requires a relatively complicated dual-chamber configuration that is separated by an appropriate ion exchange membrane, and contains anode, cathode and well-separated anolyte and catholyte. In addition, the bulky carbon-based electrodes in conventional MFCs are not compatible with the microfabrication processes described above. Nevertheless, several groups have designed intricate microscale chambers for performing MFC research at this new dimension (Table 1). A 10-ml single-chamber reactor has been constructed using a Gortex gasket to study the electrical coupling between Shewanella putrefaciens and a surface-modified gold anode [9]. The authors have observed an extremely low background current of 0.1 nA and an instant start-up (i.e. the time interval between bacterial inoculation and the onset of current generation), which has enabled them to distinguish fine-scale changes in current generation as a function of progressive anode surface modification. In addition, milliliter-MFC arrays that are individually addressable are emerging as a versatile platform for highthroughput identification and characterization of electrochemically active microbes. Compared with conventional H-shaped, two-chamber MFCs, milliliter-MFC arrays consume less materials and reagents, enable parallel, repro-
Review Box 1. Microfluidics for manipulating the bacterial microenvironment Incubation of bacterial cells generally involves the control of a series of physical, chemical and biochemical conditions, including temperature, osmotic pressure, pH, electrochemical potential, illumination, nutrient/gas supply, removal of metabolic waste, and exclusion of contamination [42]. Conventional flask or agar-plate-based culture approaches can provide appropriate conditions for growth of large populations of cells, yet use of these methods can be difficult (and sometimes impossible) when precision of control in the microenvironment around individual cells is required. In this regard, microfluidic technologies, which offer a powerful tool to manipulate fluids in a predicable manner at micrometer dimensions [43], can provide unprecedented control of the cellular microenvironment. Microfluidic systems are typically composed of well-defined, micro- to nanoliter channels or chambers on a surface. These are not simply scaled-down versions of macrofluidic devices, because fluid behavior changes significantly when the channel/ chamber size is reduced to a critical value (tens to hundreds of microns depending on specific conditions) [44,45]. Uniform chemical conditions are maintained in each flow layer, while mass transport between the layers is diffusion-limited. Fluid dynamics in microfluidic systems can be predetermined by patterned channels and specially designed micro-components (e.g. valves and pumps), and therefore, is highly predictable and controllable. Microfluidic systems can be constructed by well-established microfabrication processes, as exemplified by photolithography. In brief, photolithography is used to transfer a computer-designed pattern from a mask to a substrate mediated by photosensitive polymer (PR). The topographically patterned substrate can subsequently be used as templates for preparing a variety of microfluidic systems. Figure 2 briefly outlines the technical procedures in fabrication of three representative types of microfluidic chambers, made of silicon, PR and PDMS, respectively.
ducible and cost-effective analysis of electrogenic strains, and thus can greatly accelerate MFC research. For example, a 24-well MFC array has been developed using gold as anode material [47,48]. Each well has an anode chamber volume of about 650 ml and functions as an independent MFC. The MFC array prototype has been used to screen environmental microbes. Using this array, researchers have succeeded in isolating a bacterial strain that displays 2.3-fold higher power output than the Shewanella oneidensis MR-1 reference strain. In another case, multi-anode/ common cathode MFC arrays have been created by employing commercially available 1-ml pipette tips as MFC chambers, each with an anode chamber volume of 400 ml [49]. The resultant array of micro-MFCs allows for efficient monitoring of bacterial growth cycles and comparison of carbon source utilization of S. oneidensis MR-1 species. Besides the micro-chambered MFCs with fixed-volume fluids, micro-MFCs that contain microfluidic channels that allow fluid exchange have also been developed. The earliest report was a silicon-based microfluidic MFC system [50]. Two silicon wafers were first micro-machined to embed convoluted microfluidic channels, followed by wet-etch to form through-holes that served as fluidic inlets and outlets. Using an evaporated gold layer as electrodes, two wafers were pressed face-to-face, yet were separated by a PEM film. Glucose was used as the carbon source. Relative to the negative control, inoculation of Saccharomyces cerevisiae in this 16-ml MFC yielded an apparent current of 15 mA/ cm2 (on a 10-V load for 14 min), which corresponded to a
Trends in Biotechnology February 2011, Vol.29, No. 2
power output of 0.5 W/m3. Using a modified micro-MFC topped with transparent glass covers, the authors later reported current generation from a photosynthetic strain of Cyanophyta [51]. The new micro-MFC generated a current via photosynthesis when illuminated, and via glucose fermentation in the dark. Under a 10-V load, the power density was 40 pW/cm2. In a more recent study, a 15-ml polydimethylsiloxane (PDMS)-based micro-MFC with micropillar anodes has been demonstrated [52]. By using the same strain (S. cerevisiae) and a 1-kV load, the optimized device produced an increased current of 4.3 mA/ cm2, and a power of 42.4 nW/cm2 (3.4 W/m3, as calculated from the reported data), with output sustained for 60 min. Micro-MFC case study: Shewanella Here, we take our recently developed 1.5-ml MFC device as an example to discuss the fabrication and measurement of micro-MFCs [53]. This dual-chamber microfluidic MFC allows on-chip bacterial culture and conversion of bacterial metabolism into electricity, by utilization of the smallest anode chamber for all reported MFCs. As shown in Figure 3a and b, the micro-MFC was constructed by sequentially stacking a silicon wafer prefabricated with a gold-film anode and an SU-8 microfluidic channel (1.5 ml), a PEM, a carbon-cloth cathode, and a PDMS cathode chamber (4 ml). The device assembly was sandwiched between two acrylic plates. The well-defined channel thicknesses (100 mm) provided a short distance between the electrodes, thereby allowing efficient proton diffusion while reducing the electrolyte resistance. Inoculation of S. oneidensis MR-1 in the MFC anode chamber yielded a pronounced current density of 1.3 mA/cm2 on a 100-V resistor, with current sustained for 20 h. The current generation was reproducible when operated in a batchfed mode (Figure 3c). The Coulombic efficiency of the MFC that consumed lactate was calculated to be 2.8% – much lower than macroscopic MFCs (up to 90% [54]) – which indicates that the chemical-to-electricity conversion has not yet been optimized and represents an area that requires improvement. Electron microscopy images have shown that the MR-1 cells grow on the gold anode with high density and in a multi-layered arrangement (Figure 3c, inset). Remarkably, comparative studies of bacterial growth in various reactors have revealed an enhanced cell density on the gold anode in a 1.5-ml micro-MFC (>2 106 cell/mm2) compared to that in a 5-ml Petri dish (5 104 cell/mm2). These results were consistent with the increased current density and short start-up phase of micro-MFCs [9,50–53], which suggests that cell-to-anode coupling was enhanced in the micro-chamber environments. Polarization and power curves have revealed that the micro-MFC (Figure 3d) operates in a region where Ohmic loss was dominant [55] and the maximal power generated was 23 nW (15.3 W/m3). Linear fitting of the polarization curve yielded a relatively high internal resistance of 13 kV, which is orders of magnitude higher than that of the optimized milliliter-MFCs (down to several V) [23,56]. The high internal resistance could have stemmed from the small electrode area, polarization loss owing to slow electrode reactions, and/or electrolyte/PEM resistances 65
()TD$FIG][ Review
Trends in Biotechnology February 2011, Vol.29, No. 2
(a)
K3Fe(CN)6
(b) Catholyte 1
PDMS
2
Carbon cloth PEM
Anolyte
Growth medium
R
+ SU-8
3
C MF
Gold
ro-
Mic
(c)
Si wafer
2 cm
(d)
2.0
50
25
40
20
30
15
20
10
10
5
0
0
Voltage (mV)
1.0
Power (nW)
Current (µA)
1.5
Lactate
0.5
0.0 0
20
40
60
80
100 120 140 160 180
Time (h)
0
0.5
1.0
1.5
2.0
Current (µA) TRENDS in Biotechnology
Figure 3. Fabrication and performance of a 1.5-ml MFC device. (a) Schematic presentation of the MFC design and the key components. (b) Photograph of the microfluidic MFC device described in (a), filled with electrolytes. (c) Current recorded from a batch-fed micro-MFC over time. The micro-MFC generated current in response to 20 mM lactate addition (indicated by arrows) and sustained the current for 20–30 h. Inset: scanning electron microscopy image of the bacteria colony at the anode. Scale bar, 2 mm. (d) Polarization (solid symbols) and power (empty symbols) curves measured from the micro-MFC device. Reproduced with permission from Ref. [53].
that scaled inversely with the electrode area. The observed high internal resistance limits power output and represents a major challenge for all (sub)microliter-scale MFCs [56]. To solve this problem, the development of new electrode materials [25,27,57] with high surface area, low resistance, and strong affinity for bacterial cells (facilitating extracellular electron transfer) is crucial. To date, the potential of utilizing microfluidic devices for MFC research has not been fully explored, although such devices have demonstrated several general advantages over conventional MFC devices. As a result of the size reduction, micro-MFCs show low background signals, rapid start-up, and high sensitivity to bacterial metabolism. Most importantly, the much smaller electrode in the microMFC is more responsive to a local electrochemical change in its microchamber than a macroscopic device, which might allow MFC research to enter the realm of measurement of electrogenesis by a single bacterial cell. Key challenges for miniaturized MFCs The development of micro-MFCs offers new and unique opportunities to analyze MFC reactions at small scales in a well-controlled microenvironment. However, miniaturization of the electrodes and the reaction chamber also present 66
challenges for micro-MFCs. One of the major challenges is to reduce their large internal resistance, which, if it can be solved, could lead to better output power densities and potential applications as power supplies for ultra-small electronics. Theoretically, the membrane and electrolyte resistances can be reduced by carefully designed reactor architectures; for example, by designing chambers with even higher SAV ratios, using a thinner PEM, and increasing the ionic concentration of electrolytes to the limits tolerated by the bacterial cells [55]. Systematic exploration of growth conditions and alternative strains could help to identify the best bacterial candidates for proliferation and electrogenesis in the microfluidic environment that can differ significantly from that of conventional cultivation conditions. Towards reducing resistance associated with the electrodes, incorporation of high-surface-area, carbon-based electrodes in the microscale chambers might offer a promising solution. Our recent result of a 4-ml carbon clothanode MFC exhibited enhanced power output (250 nW) and power density (62.5 W/m3) compared to the 1.5-ml goldanode MFC (23 nW, 15 W/m3) operated under identical conditions (D. Morse, unpublished). It is desirable to explore aggressively non-conventional nanoscale electrodes, such as carbon nanotubes, graphene or semiconductor
Review nanowires, for use in micro-MFCs. Additionally, one also must pay attention to the design of the cathode half-cells to prevent cathode limitation. Another key scientific challenge is the coupling of fundamental electrochemical studies to the microfluidic MFC platform. Although rational design of micro-reactors can lead to an optimized solution for a micro-MFC with good power densities, analytical electrochemical measurements provide complementary information for a quantitative understanding of the electrode kinetics, bacterial redox potential, mass transport, and internal resistance distribution. However, most important electrochemical measurements presently require the use of a bulky reference electrode, which cannot yet be coupled into a microscale chamber. Despite the fact that planar Ag/AgCl microelectrodes have been demonstrated as reference electrodes elsewhere, their stabilities and lifetimes are still insufficient to support long-term MFC studies [58]. Therefore, the development of a reliable three-electrode microelectrochemical system would be crucial for enabling a series of fundamental electrochemical characterizations in microfluidic MFCs. Future prospects Compared to the well-established milli- and macro-MFCs, micro-MFCs are still in their infancy. Although the small power output and high fabrication cost exclude microMFCs from large-scale use in power supply and waterpurification applications, their unique dimensions provide advantages for microfluidic MFCs in several novel areas. Ultra compact and fast current-based assays High-density integration of functional micro-MFCs could allow researchers to conduct multiple electrical, electrochemical or biochemical measurements in parallel. Considering a micro-MFC array composed of MFC units with a side-length of 100 mm and a spacing of 100 mm, 2500 individually operated MFCs can be arrayed within a 1cm2 area. As discussed above, high-SAV-ratio microscale reactors can shorten the ion diffusion length and promote biofilm development to convert analytes (substrates) more efficiently into detectable current signals. Such arrays of microliter-volume MFCs thus offer unique advantages of fast start-up, high sensitivity and superior microfluidic control over the measured microenvironment, which makes them good candidates for rapid screening of electrode materials, bacterial strains and growth media. Thus, for instance, the micro-MFC array can be used to select optimally efficient electrogenic bacterial strains, which can then be used in conventional MFC reactors on larger scales. Bioenergy sources for ultra-small electronics With the development of nanoscience and technology, a broad range of nanodevices have been demonstrated for the next-generation applications in computing, communication and sensing [59,60]. These nanoscale electronic elements and networks generate, process and transmit signals on a low-energy basis (nW to mW) [59], which can in principle be supplied by micro-MFCs. Through on-chip integration, the combination of microbial energy
Trends in Biotechnology February 2011, Vol.29, No. 2
supply and nanodevice operation could lead to self-sustained nanosystems that might find significant uses in diverse areas. Further studies are still needed to test whether the long-term operation of micro-MFCs is practical, and how to overcome potential challenges, such as cells clogging the microchannel. Real-time visualization of electrogenic cells in an MFC The microscale thickness of micro-MFCs is compatible with the working distances of conventional optical microscopes and thus can be directly placed under an optical microscope for observing live bacterial behavior. The ability to visualize live cells in situ in an MFC requires device transparency for light transmission, which can be easily obtained by PDMS- or glass-based micro-MFCs. In addition, an enhanced optical contrast between cells and the environment is necessary for live-cell investigation, which can be achieved using staining dyes [61], genetically modified fluorescent strains [61,62] or phase-contrast imaging [63]; albeit, these requirements might add to the complexity of the reactor. Correlation of optical and electrical/ electrochemical studies of MFCs could provide complementary information on cell growth and functioning in a microMFC, and this in turn could suggest novel strategies for unravelling and optimizing the details of extracellular electron transfer mechanism in MFCs. Single-bacterium studies One of the most attractive promises of micro-MFCs is to study a single bacterium, which could revolutionize our understanding in these electricity-generating microorganisms. To date, the MFC field has largely focused on the collaborative performance of a large population of bacterial cells; typically in the form of a biofilm. The biofilm is a complex mixture of dead, live and dividing cells, embedded within a self-produced matrix of extracellular polymeric substances [64,65]. Mass transport and electron transfer in the biofilm is unknown, complex and uncontrolled. Measurements of the electrogenic performance of biofilms yield averaged results from diverse and changing populations, which makes it difficult to achieve reproducibility and to obtain precise, quantitative information on specific events in the microbial electron transfer chain. By contrast, single-bacterium analyses could allow us to elucidate this basic information from individual bacterial cells in a controlled microenvironment, which could provide a higher-resolution perspective in understanding the key electrochemical and biochemical processes at the bacterial/ electrode interface. Only with further in-depth comprehension of these fundamental processes will we be able to improve the efficiency of bacterial electrogenesis and further advance MFC research and development. Figure 4 illustrates a possible design of a single-bacterium microfluidic MFC device. Microelectrodes are first evaporated onto the substrate and coupled in the microfluidic channel. These microelectrodes are individually addressable, serve as anodes, and share a common PEC/ cathode assembly, thus forming a microelectrode MFC array. On this unique platform, the maximal number of bacterial cells coupled to the anode can be controlled by individual anode size. Intercellular communication among 67
()TD$FIG][ Review
Trends in Biotechnology February 2011, Vol.29, No. 2
Microfluidic channel
Fluid out
Fluid in
Microelectrode
TRENDS in Biotechnology
Figure 4. Artistic rendering of a single-bacterium micro-MFC device. Microelectrode arrays are defined within the microfluidic anode chamber. By control of the electrode size, spacing and cell density, such a device should make it possible to probe electricity generation from a single electrogenic cell.
bacterial cells (e.g. quorum sensing) could be studied as a function of cell density with such a device. The ability to create complex electrode patterns with well-defined morphologies (e.g. pillars, trenches and holes), combined with the precise control of electrode position, could be used to tune the 3D geometry of the microbial/anode interface at the (sub)microscale. Moreover, the incorporation of nanoscale electrodes, such as nanowire arrays, in the microfluidic platform might enable multi-point, intracellular measurements of a single bacterium, which introduces further exciting opportunities in fundamental MFC research. Acknowledgements We thank Dr. Ning Cao at the University of California at Santa Barbara Nanofabrication Center for valuable discussions. This work was supported in part by grants from the U.S. Department of Energy (grant # DE-FG02-02ER46006) and the U.S. Army Research Office (through contract no. W911NF-09-D-0001 to the Institute for Collaborative Biotechnologies). F. Q. thanks the support in part of this work by the National Science Foundation (award # CBET 1034222).
References 1 Bond, D.R. et al. (2002) Electrode-reducing microorganisms harvesting energy from marine sediments. Science 295, 483–495 2 Lovley, D.R. (2006) Bug juice: harvesting electricity with microorganisms. Nat. Rev. Microbiol. 4, 497–508 3 Logan, B.E. et al. (2006) Microbial fuel cells: methodology and technology. Environ. Sci. Technol. 40, 5181–5192 4 Logan, B.E. (2008) Microbial Fuel Cells, John Wiley 5 Fredrickson, J.K. et al. (2008) Towards environmental systems biology of Shewanella. Nat. Rev. Microbiol. 6, 592–603 6 Bretschger, O. et al. (2007) Current production and metal oxide reduction by Shewanella oneidensis MR-1 wild type and mutants. Appl. Environ. Microbiol. 73, 7003–7012 7 Chaudhuri, S.K. and Lovley, D.R. (2003) Electricity generation by direct oxidation of glucose in mediatorless microbial fuel cells. Nat. Biotechnol. 21, 1229–1232 8 Bond, D.R. and Lovley, D.R. (2003) Electricity production by Geobacter sulfurreducens attached to gold electrodes. Appl. Environ. Microbiol. 69, 1548–1555 9 Crittenden, S.R. et al. (2006) Mediating electron transfer from bacteria to a gold electrode via a self-assembled monolayer. Langmuir 22, 9473–9476 10 Newman, D.K. and Kolter, R. (2000) A role for excreted quinones in extracellular electron transfer. Nature 405, 94–97
68
11 Marsili, E. et al. (2008) Shewanella secretes flavins that mediate extracellular electron transfer. Proc. Natl. Acad. Sci. U. S. A. 105, 3968–3973 12 Reguera, G. et al. (2005) Extracellular electron transfer via microbial nanowires. Nature 435, 1098–1101 13 Gorby, Y.A. et al. (2006) Electrically conductive bacterial nanowires produced by Shewanella oneidensis strain MR-1 and other microorganisms. Proc. Natl. Acad. Sci. U. S. A. 103, 11358–11363 14 Reguera, G. et al. (2006) Biofilm and nanowire production leads to increased current in Geobacter sulfurreducens fuel cells. Appl. Environ. Microbiol. 72, 7345–7348 15 Pessanha, M. et al. (2004) Redox characterization of Geobacter sulfurreducens cytochrome c7: physiological relevance of the conserved residue F15 probed by site-specific mutagenesis. Biochemistry 43, 9909–9917 16 Rabaey, K. et al. (2007) Microbial ecology meets electrochemistry: electricity-driven and driving communities. Int. Soc. Microb. Ecol. J. 1, 9–18 17 Aelterman, P. et al. (2008) The anode potential regulates bacterial activity in microbial fuel cells. Appl. Microbiol. Biotechnol. 78, 409–418 18 Firer-Sherwood, M. et al. (2008) Electrochemical interrogations of the Mtr cytochromes from Shewanella: opening a potential window. J. Biol. Inorg. Chem. 3, 849–854 19 He, Z. and Mansfeld, F. (2009) Exploring the use of electrochemical impedance spectroscopy in microbial fuel cell studies. Energy Environ. Sci. 2, 215–219 20 Liu, H. et al. (2004) Production of electricity during wastewater treatment using a single chamber microbial fuel cell. Environ. Sci. Technol. 38, 2281–2285 21 Cheng, S. et al. (2006) Increased power generation in a continuous flow MFC with advective flow through the porous anode and reduced electrode spacing. Environ. Sci. Technol. 40, 2426–2432 22 Ringeisen, B.R. et al. (2006) High power density from a miniature microbial fuel cell using Shewanella oneidensis DSP10. Environ. Sci. Technol. 40, 2629–2634 23 He, Z. et al. (2006) The upflow microbial fuel cell with an interior cathode: assessment of the internal resistance by impedance spectroscopy. Environ. Sci. Technol. 40, 5212–5217 24 Fan, Y.Z. et al. (2007) Enhanced Coulombic efficiency and power density of air-cathode microbial fuel cells with an improved cell configuration. J. Power Sources 171, 348–354 25 Dumitru, A. et al. (2008) Biofilm growth from wastewater on MWNTs and carbon aerogels. Physica. Status Solidi (a) 205, 1484–1487 26 Richter, H. et al. (2008) Electricity generation by Geobacter sulfurreducens attached to gold electrodes. Langmuir 24, 4376–4379 27 Qiao, Y. et al. (2008) Nanostructured polyanifine/titanium dioxide composite anode for microbial fuel cells. ACS Nano. 2, 113–119 28 Wang, X. et al. (2009) Use of carbon mesh anodes and the effect of different pretreatment methods on power production in microbial fuel cells. Environ. Sci. Technol. 43, 6870–6874 29 Liu, Y. et al. (2010) The study of electrochemically active biofilms on different carbon-based materials in microbial fuel cells. Biosens. Bioelectron. 25, 2167–2171 30 Rosenbaum, M. et al. (2010) Light energy to bioelectricity: photosynthetic microbial fuel cells. Curr. Opin. Biotechnol. 21, 259–264 31 Izallalen, M. et al. (2008) Geobacter sulfurreducens strain engineered for increased rates of respiration. Metab. Eng. 10, 267–275 32 Xing, D.F. et al. (2008) Electricity generation by Rhodopseudomonas palustris DX-1. Environ. Sci. Technol. 42, 4146–4151 33 Yi, H. et al. (2009) Selection of a variant of Geobacter sulfurreducens with enhanced capacity for current production in microbial fuel cells. Biosens. Bioelectron. 24, 3498–3503 34 Gil, G.C. et al. (2003) Energy from algae using microbial fuel cells. Biotechnol. Bioeng. 103, 1068–1076 35 Min, B. and Logan, B.E. (2004) Continuous electricity generation from domestic wastewater and organic substrates in a flat plate microbial fuel cell. Environ. Sci. Technol. 38, 5809–5814 36 Selembo, P.A. et al. (2009) High hydrogen production from glycerol or glucose by electrohydrogenesis using microbial electrolysis cells. Int. J. Hydrogen Energy 34, 5373–5381 37 Lalaurette, E. et al. (2009) Hydrogen production from cellulose in a twostage process combining fermentation and electrohydrogenesis. Int. J. Hydrogen Energy 34, 6201–6210
Review 38 Dekke, A. et al. (2009) Analysis and improvement of a scaled-up and stacked microbial fuel cell. Environ. Sci. Technol. 43, 9038–9042 39 He, Z. et al. (2005) Electricity generation from artificial wastewater using an upflow microbial fuel cell. Environ. Sci. Technol. 39, 5262– 5267 40 Cheng, S. and Logan, B.E. (2007) Sustainable and efficient biohydrogen production via electrohydrogenesis. Proc. Natl. Acad. Sci. U. S. A. 104, 18871–18873 41 Cao, X. et al. (2009) A new method for water desalination using microbial desalination cells. Environ. Sci. Technol. 43, 7148–7152 42 Wilkinson, J.F. (1972) Introduction to Microbiology, Blackwell Scientific 43 Whitesides, G.M. (2006) The origins and the future of microfluidics. Nature 442, 368–373 44 Atencia, J. and Beebe, D.J. (2005) Controlled microfluidic interfaces. Nature 437, 648–655 45 Whitesides, G.M. et al. (2001) Soft lithography in biology and biochemistry. Annu. Rev. Biomed. Eng. 3, 335–373 46 Weibel, D.B. et al. (2009) Microfabrication meets microbiology. Nat. Rev. Microbiol. 5, 209–218 47 Hou, H. et al. (2009) Microfabricated microbial fuel cell arrays reveal electrochemically active microbes. PLoS ONE 4, e6570 48 Hou, H. et al. (2011) Air-cathode microbial fuel cell array: a device for identifying and characterizing electrochemically active microbes. Biosens. Bioelectron. 26, 2680–2684 49 Biffinger, J.C. et al. (2009) Characterization of electrochemically active bacteria utilizing a high-throughput voltage-based screening assay. Biotechnol. Bioeng. 102, 436–444 50 Chiao, M. et al. (2003) Micromachined microbial fuel cells, In Proceedings of 16th IEEE MicroElectro Mechanical Systems Conference, Kyoto, Japan. pp. 383–386 51 Chiao, M. et al. (2006) Micromachined microbial and photosynthetic fuel cells. J. Micromech. Microeng. 16, 2547–2553
Trends in Biotechnology February 2011, Vol.29, No. 2 52 Siu, C. and Chiao, M. (2008) A microfabricated PDMS microbial fuel cell. J. Micro Electrochem. Syst. 17, 1329–1341 53 Qian, F. et al. (2009) A 1.5-microliter microbial fuel cell for on-chip bioelectricity generation. Lab. Chip. 9, 3076–3081 54 Chang, I.S. et al. (2004) Continuous determination of biochemical oxygen demand using microbial fuel cell type biosensor. Biosens. Bioelectron. 19, 607–613 55 Fan, Y. et al. (2008) Quantification of the internal resistance distribution of microbial fuel cells. Environ. Sci. Technol. 42, 8101–8107 56 Logan, B.E. and Regan, J.M. (2006) Microbial fuel cells – challenges and applications. Environ. Sci. Technol. 40, 5172–5180 57 Peng, L. et al. (2010) Carbon nanotubes as electrode modifier promoting direct electron transfer from Shewanella oneidensis. Biosens. Bioelectron. 25, 1248–1251 58 Polk, B.J. et al. (2006) Ag/AgCl microelectrodes with improved stability for microfluidics. Sens. Actuators B Chem. 114, 239–247 59 Li, Y. et al. (2006) Nanowire electronics and optoelectronic devices. Mater. Today 9, 18–27 60 Lieber, C.M. and Wang, Z.L. (2007) Functional nanowires. MRS Bull. 32, 99–104 61 Franks, A.E. et al. (2010) Microtoming coupled to microarray analysis to evaluate the spatial metabolic status of Geobacter sulfurreducens biofilms. Int. Soc. Microb. Ecol. J. 4, 509–519 62 Teal, T.K. et al. (2006) Spatiometabolic stratification of Shewanella oneidensis biofilms. Appl. Microbiol. Biotechnol. 72, 7324–7330 63 Jiang, X. et al. (2010) Probing electron transfer mechanisms in Shewanella oneidensis MR-1 using a nanoelectrodes platform and single-cell imaging. Proc. Natl. Acad. Sci. U. S. A. 107, 16806–16810 64 Dunne Jr, W.M. (2002) Bacterial adhesion: seen any good biofilms lately. Clin. Microbiol. Rev. 15, 155–166 65 Biffinger, J.C. et al. (2007) A biofilm enhanced miniature microbial fuel cell using Shewanella oneidensis DSP10 and oxygen reduction cathodes. Biosens. Bioelectron. 22, 1672–1679
69