A Micro Fabricated PDMS Microbial Fuel Cell 2008

JOURNAL OF MICROELECTROMECHANICAL SYSTEMS, VOL. 17, NO. 6, DECEMBER 2008 1329

A Microfabricated PDMS Microbial Fuel CellChin-Pang-Billy Siu, Student Member, IEEE, and Mu Chiao, Member, IEEE

Abstract—This paper presents a microfabricated polydimethyl-siloxane (PDMS) microbial fuel cell (MFC) with embeddedmicropillar electrodes. This MFC is characterized by a flexi-ble and biocompatible structure suitable for body implantationas a potential power source for implanted bioMEMS devices.The MFC is biocatalyzed by a microorganism, Saccharomycescerevisiae, which converts chemical energy stored in glucose inthe blood stream to electrical energy. The MFC is a laminate de-sign, consisting of 0.2-μm-thick gold-evaporated PDMS anode andcathode separated by a Nafion 117 proton exchange membrane.These electrode surfaces feature more than 70 000 8-μm-highmicropillar structures in a 1.2-cm × 1.0-cm geometric area.The MFC is encapsulated by PDMS and has an overall size of1.7 cm × 1.7 cm × 0.2 cm and a net weight of less than 0.5 g.Compared with recent silicon micromachined MFCs working in aphosphate buffer medium, the presented MFC with its micropillarstructure showed a 4.9 times increase in average current densityand a 40.5 times increase in average power density when operatedat identical conditions. Using 15-μL droplet of human plasma,containing 4.2-mM blood glucose, the PDMS MFC demonstrateda maximum open circuit potential (OCP) of 488.1 mV, maximumcurrent density of 30.2 μA/cm2, and a maximum power density of401.2 nW/cm2. When the MFC operated continuously for 60 min,it showed an average OCP of 297.4 mV, average current density at4.3 μA/cm2, and average power density of 42.4 nW/cm2 at 1-kΩload. The coulombic efficiency of electron conversion from bloodglucose was 14.7%. [2008-0039]

Index Terms—Biofuel cell, blood glucose, human plasma, mi-crobial fuel cell (MFC), microfabrication, micropillar electrode,polydimethylsiloxane (PDMS).

I. INTRODUCTION

IMPLANTABLE BioMEMS devices provide new medicaltherapies for patients suffering from chronic diseases and

functional disorders [1]–[14]. For example, deep brain stimula-tors are used to treat Parkinson’s disease [2]–[4], and retinalmicroimplants have been developed for restoring the visionto patients suffering from retinitis pigmentosa and maculardegeneration [4]–[9]. Cochlear microdevices are used to reducehearing impairment [10]–[12], and intraspinal microelectrodeshave been proposed to stimulate the muscle response in casesof spinal cord injury [9], [13], [14]. These devices use electricalstimulation on nerve cells to evoke organ functions [2]–[14].The neural activation threshold generally requires 50 mV, witha 1-ms pulsewidth for signal transmission [15]. Conventional

Manuscript received February 15, 2008; revised August 15, 2008. Currentversion published December 4, 2008. This work was supported by Sweet PowerInc. Subject Editor A. J. Ricco.

The authors are with the Department of Mechanical Engineering, TheUniversity of British Columbia, Vancouver, BC V6T 1Z4, Canada (e-mail:[email protected]).

Color versions of one or more of the figures in this paper are available onlineat http://ieeexplore.ieee.org.

Digital Object Identifier 10.1109/JMEMS.2008.2006816

lithium batteries can generate 2.2 V [16]; however, their weight(12.5–15.5 g), rigid packaging, and need for surgical replace-ment may restrict their use as implants in soft tissues and organs[1], [16]–[18]. Therefore, an alternative power source is neededfor implantable microdevices [18]–[20].

The use of microbial fuel cells (MFCs) and glucose togenerate electricity has been proposed as a potential powersource for implanted microdevices [20]–[23]. They convertchemical energy directly to electricity without combustion [24].In contrast to the conventional hydrogen and solid oxide fuelcells that require complex fuel reforming and sophisticatedstorage solutions [25]–[27], glucose fuel is available in theblood stream and is replenished from food. A stable glucoseconcentration in the blood ranges between 0.7 g/L (3.9 mM)and 0.9 g/L (5.0 mM) and is regulated by insulin secretionfrom the pancreas [28]. In addition, other body fluids, such assubcutaneous fluids and cerebrospinal fluid, have glucose con-centrations ranging from 2.2 to 25 mM [29]–[31]. Moreover,MFCs have some distinctive advantages. For example, theyuse microbia as the self-replicating catalysts to oxidize organicand inorganic matter at mild temperatures (20 ◦C–40 ◦C) andnearly neutral acidity [32]–[34]. In comparison with enzymicbiofuel cells, which usually adopt a particular enzyme as thecatalyst, MFCs use multiple endogenous enzymes to effectivelydissociate the fuel into simple molecules for complete energyextraction [35]. Moreover, MFCs require no specific enzymeisolation, purification, and stabilization on the electrode surface[21], [33], [35].

The electrical effects associated with microbial metabolismwere initially studied in 1911. An open circuit potential (OCP)of 0.32 V was reported from 5 g of Saccharomyces cerevisiae(S. cerevisiae) mixed with a 0.55-M glucose culture medium[36]. In 1931, an electric potential of 35 V was generated froma batch of MFCs [37]. Microorganisms such as A. eutrophus,A. nidulans, C. butyricum, C. diphtherice, E. coli, M. cereficans,M. lysodeikticus, P. Vulgaris, and S. cerevisiae were used inMFC research [21], [33]. Early studies led to a limited currentdensity of about 10 μA/cm2, regardless of the volume of cul-tured electrolyte or the electrode surface area [38]. One of themajor factors accounting for this limitation is the nonconductivenature of the cytoplasmic membrane, which inhibits the trans-mission of electrons from the microbial interior to the electrode[39], [40]. An increase in current output was achieved by theaddition of electron mediators to the anodic culture medium[40]–[44].

Electron mediators are low molecular weight compoundsfrom phenoxazine, phenothiazine, phenazine, indophenol,or bipyridilium groups [23], [45], [46]. The molecularweight of electron mediators for MFC should be less than600 Da to be able to penetrate through the porous microbial cell

1057-7157/$25.00 © 2008 IEEE

Authorized licensed use limited to: CINVESTAV IPN. Downloaded on May 17,2010 at 15:12:20 UTC from IEEE Xplore. Restrictions apply.

1330 JOURNAL OF MICROELECTROMECHANICAL SYSTEMS, VOL. 17, NO. 6, DECEMBER 2008

wall of yeast [47]. Electron mediators can reduce the metaboliccoenzymes and shuttle the endogenous electrons and protons tothe electrode of MFCs [46], [48]. The electron-harvesting per-formance of electron mediators depends on their redox rate withcoenzymes and electrodes, redox potential, chemical stability,affinity with the microbia, solubility in the medium as well asthe pH, and temperature of the medium [21], [40]. Commonelectron mediators adopted for use in MFCs include thionine,methyl viologen, 2-hydroxy-1, 4-napthoquinone, neutral red,and methylene blue (MB) [24]. For example, E. coli. MFCshave shown a current increase from 0.5 to 2 mA after 25-mgMB was added to the 400-mL anodic culture medium [38].Some electron mediators (e.g., MB) have been discovered tohave therapeutic applications [49]–[51], although their muta-genic and other unintended biological effects must be fullyanalyzed and neutralized before they can be applied to thehuman body [52], [53]. This precaution also extends to potas-sium ferricyanide which is a common oxidant in the cathodecompartment of MFCs [24], [33], [41], [54].

Recently, metal-reducing geobacteria, such as Geobactermetallireducens, Rhodoferax ferrireducens, and Shewanella pu-trefaciens, have been used as biocatalysts [35], [55], [56].These species have conductive enzymes, crytochromes, in theirouter cellulous membrane that provide an electron conductionpathway from the intercellular cell body to the electrode surfacewithout needing an electron mediator [34], [57]. Althoughmediator-less MFCs have a coulombic efficiency of over 80%,some of these microorganisms can catalyze glucose only whenmetallic ions such as Fe3+ ions are present in the anodic me-dium as the primary metabolic substrate [55]. Thus, the focusis on using them in external power applications [32], [35], [37].

Most MFCs were developed in macroscale, with 10–400 mLvolume per electrode compartment [23], [34], [36], [38], [40]–[44], [55], [56]. Some of these MFCs use continuous stirring,nitrogen or argon bubbling, acidity control, and computerizednutrient feed mechanism to control the reaction rate. In addi-tion, they are assembled with porous, reticulated, mesh, graph-ite felt, or platinum electrodes to optimize performance [21],[27], [35], [44], [55]. Recently, miniaturized MFCs are exploredthrough microfabrication [58], [59]. One of the challenges inminiaturization is to increase current and power density withina limited electrode surface area and restricted compartmentcavity size [35]. Microfabrication provides batch productionand structure uniformity [1]. Tailor-made microstructures canbe fabricated on miniaturized electrodes to increase the reactionsurface area and provide a favorable scaffold for microbialgrowth [1], [60]–[63]. The silicon micromachined MFC em-ployed a microchannel design and achieved 1.3-μA/cm2 av-erage current density and 0.2-nW/cm2 average power densityat 101-Ω load for 40 min. It used 16 μL of 0.1-M pH 7.0phosphate buffer with S. cerevisiae, 10-mM MB, and 1-Mglucose as the anolyte, and the same volume of phosphate bufferwith 20-mM potassium ferri(III)cyanide as the catholyte [58].

Polydimethylsiloxane (PDMS) is a flexible silicone-basedmaterial [64], [65]. It can bond with other surfaces to form wa-tertight seals [66]–[68]. Biocompatibility, hemocompatibility,and inflammatory tests have established PDMS as a potentialpolymer for in vivo applications [69], [70]. Several studies on

Fig. 1. Operating principle of the PDMS MFC.

chemical micro fuel cells have already examined PDMS as astructure material but with no surface modification [71]–[73].In this paper, we present a microfabricated PDMS MFC withmicropillar embedded on the electrode surfaces. The objectivesare as follows: 1) to evaluate the improvement of electricity out-put from the addition of a micropillar structure on the electrodeand 2) to investigate the feasibility of using human plasma forthe electricity generation. The morphological adaptation of themicroorganism on the gold-evaporated PDMS electrode is qual-itatively inspected, and the performances of the PDMS MFCsare compared with previous microfabricated MFCs. TransientOCP and current densities are reported.

II. OPERATING PRINCIPLE

Fig. 1 shows the operating principle of a PDMS MFC. Theanode and cathode compartments are separated by a protonexchange membrane. The biocatalyst, S. cerevisiae, is encap-sulated inside the anode compartment and uses glucose as fuel.The metabolic process of S. cerevisiae is a series of endoge-nous enzymatic redox reactions. Glucose is first converted topyruvate through the glycolytic pathway and then catabolizedinto simple molecules, including carbon dioxide and water, viathe tricarboxylic acid cycle. During the multienzymic process,chemical energy is captured throughout the electron transportchain. Nicotinamide adenine dinucleotide (NAD+) and nicoti-namide adenine dinucleotide dehydrogenase (NADH) functionas the coenzymes for the reactions and are repeatedly oxidizedand reduced to prompt the synthesis of the biological energyunit, adenosine triphosphate (ATP) [71], [74]. MB is used as anelectron mediator, which can penetrate to the microbial interiorand reduce the electrons from the oxidation between NADHand NAD+ [23]. These endogenous reactions are as follows[75]–[77]:

C6H12O6 + 6O2 → 6CO2 + 6H20 (1)NADH →NAD+ + H+ + 2e− (2)

MB + NADH →MBH + NAD+. (3)

The reduced MBH then diffuses back to the extraneousmedium and is oxidized on the electrode surface to liberate


SIU AND CHIAO: MICROFABRICATED PDMS MICROBIAL FUEL CELL 1331

Fig. 2. Exploded view of the fuel cell structure: (a) PDMS MFC assembly and (b) the PDMS electrode with micropillar structures.

protons and electrons. To maintain the intercellular pH gradi-ent, the microorganism also releases protons from metabolismto the anodic medium [78]. The proton exchange membranerestricts gas and electron from crossing over but prefers protontransmission across the two electrode compartments, whereasthe potential difference between the anode and cathode drivesthe electron flow through an external circuit. Potassium ferri-cyanide is used as an oxidant in the cathode compartment withthe advantage of no leakage transport through the Nafion protonexchange membrane. Cyclic voltammetric studies showed thatthe crossover of ferricyanide/ferrocyanide ion is prohibited dueto the charge repelling from the osmotic characteristic of Nafionand a bigger molecular size than the 5-nm diameter or smallermembrane porosity [79]–[81]. Ferricyanide ion (Fe(CN)3

−6 ) in

the cathode is first reduced to ferrocyanide ion (Fe(CN)4−

6 ),and the ferrocyanide ion then reacts with a proton and oxygento oxidize back to ferricyanide ion with the production of water(H2O). The electrochemical reactions and the standard equi-librium potential (versus normal hydrogen electrode at 25 ◦C,1 atm, and 1-M concentration) for the anode and cathode aredescribed as [41], [82], [83] follows:Anode

MBH → MB + H+ + 2e− E0anode = −0.145 V (4)

Cathode

4Fe(CN)3−6 + 4e− → 4Fe(CN)4−6 (5)

4Fe(CN)4−6 + 4H+ + O2 → 4Fe(CN)3−6 + 2H20 (6)

Combined (5) and (6)

O2 + 4H+ + 4e− → 2H20 E0cathode = +0.82 V (7)

Overall redox reaction

2MBH + O2 + 2H+ → 2MB + 2H20. (8)

III. DESIGN AND FABRICATION

Fig. 2(a) shows an exploded view of the PDMS MFC. Thelaminate MFC structure consists of five layers. Each electrodecompartment is constructed by the insertion of a 100-μm-thickPDMS gasket in between the electrode and the proton exchangemembrane (DuPont, Nafion 117). The PDMS electrodes arefabricated with an array of micropillars in a 10-mm × 12-mmarea [Fig. 2(b)]. A thin gold coating on the PDMS surfaceserves as the conducting electrode. During experiments, fueland electrolytes are injected via one of the 500-μm-diameteraccess ports at the two corners of the PDMS electrodes. Anexternal electric connection is made via the protruding gold-evaporated surface of the electrode. In principle, the PDMSanode and cathode can be molded to form a specific mor-phology suitable for different implantation requirements [68].Other biocompatible polymeric materials such as polyimide,polylactide-co-glycolide (PLGA), polyurethane, and poly-ethylene are also suitable for this application [1], [9], [14], [68],[69], [84].

Fig. 3 shows the fabrication of the PDMS electrode using amolding process [85]. A piece of silicon wafer is first cleanedby the standard RCA-1 and RCA-2 processes [Fig. 3(a)].Then, a 5000-Å-thick silicon dioxide layer is grown on thesilicon substrate [Fig. 3(b)]. The silicon dioxide is patternedby a plasma-assisted etching process to define the geometriesof the electrode and its micropillars [Fig. 3(c)]. The siliconsubstrate is then etched by vapor phase xenon difluoride atroom temperature using 2.5-T XeF2 and 800-mT N2 partialpressures [Fig. 3(d)]. The silicon dioxide layer is removed ina buffered oxide solution (BOE 10:1) at room temperature[Fig. 3(e)]. Layers of 500-Å-thick chromium and 2000-Å-thickgold are evaporated onto the silicon surface by a DeeWongelectron beam system [Fig. 3(f)]. The gold layer serves as anintermediate layer that will later promote the PDMS demoldingprocess. A premixed 10 : 1 weight ratio PDMS elastomer andcuring agent (DuPont, Sylgard 184) is spin-coated onto the



Fig. 3. Schematic diagram of fabrication process of the PDMS micropillarelectrode. (a) Si wafer standard clean. (b) 5000-Å oxide growth. (c) PECVDoxide etch. (d) XeF2 Si dry etch. (e) BOE oxide etch. (f) 500-Å Cr/2000-ÅAu electron beam evaporation. (g) PDMS spin coating. (h) PDMS layerdetachment and fluidic access punching. (i) 500-Å Cr/2000-Å Au electron beamevaporation.

Fig. 4. Fabricated PDMS electrodes. (a) PDMS detached from a 5-Å/sgold-evaporated silicon mold, showing the fractured gold pieces delaminatedfrom the silicon on the PDMS. (b) PDMS detached from gold silicon moldwith 0.5–2-Å/s gold evaporation on the silicon surface. (c) Fabricated PDMSelectrode with gold-evaporated electrode surface.

silicon mold [Fig. 3(g)]. After 1-h degassing in a vacuumchamber, the PDMS is cured at 60 ◦C for 2 h.

The deposition rate of the Cr/Au thin films has a significanteffect on preventing separation of the film from the siliconsurface during the demolding process. Fig. 4 shows PDMSsurfaces after the demolding process. Without the intermediategold layer, the PDMS is not able to detach from the siliconwithout breaking. The silicon mold is placed 40 cm aboveand faced downward to the metal source inside the depositionchamber. Successive depositions of chromium and gold areperformed at 2.0 × 10−6 T without interruption. When the

Fig. 5. Scanning electron image of the gold-evaporated PDMS electrode, withtypical micropillar cross-sectional dimensions of 40 μm × 40 μm.

chromium and gold layers are deposited at a rate no greaterthan 2 Å/s, the peeling of the PDMS is successful with norupture of the micropillars. Therefore, replicate micromoldingcan be performed repetitively. When the deposition rate is fasterthan 2 Å/s, the PDMS can still be detached; however, part ofthe gold layer delaminates from the silicon and attaches to thePDMS surface. A similar phenomenon has been observed withchromium thicknesses of 300 and 800 Å. In any case, whetherthe delamination occurs at the Cr/Au or Cr/Si interface remainsunclear. Further studies are required.

Fluid accesses of 500-μm diameter are then mechanicallypunched at the diagonal corners of the electrode [Fig. 3(h)].Finally, 500-Å-thick chromium and 2000-Å-thick gold areevaporated onto the PDMS surface as a conducting electrode[Fig. 3(i)]. Fig. 4 shows the PDMS detached from the siliconmolds at different gold evaporation rates and the finished gold-evaporated PDMS electrode. Fig. 5 shows a scanning electronicmicroscope (SEM) picture of the micropillar geometry on thePDMS surface. A total of 74 451 micropillars are located oneach electrode. The micropillars have typical cross-sectionaldimensions of 40 μm × 40 μm and a height of 8 μm, measuredby a profilometer (Alpha Step 200, Tencor). The effectiveelectrode surface area is increased 1.8 times by the micropil-lars, changed from 1.2-cm2 flat geometric surface to 2.16-cm2

effective electrode area.Before assembling the MFC, impurities on the proton ex-

change membrane are removed by chemical treatment. First,the membrane is soaked in a boiling 1 : 9 diluted hydrogenperoxide aqueous solution for 1 h. It is then boiled in deionized(DI) water for 2 h to remove the hydrogen peroxide residue.Then, the membrane is immersed in boiling 1 : 9 sulphuricacid for 1 h and dipped in boiling DI water for another 2 hsubsequently [86]. Finally, the membrane is dried at roomtemperature. The laminate MFC structures are sealed by 10 : 1premixed PDMS. These adhesive PDMS layers, with an aver-age thickness of 10 μm, are allowed to set at room temperaturefor 24 h. The boundaries of the MFC are sealed by PDMSand set at room temperature for another 24 h. Fig. 6 showsthe assembled PDMS MFC. The overall dimensions of thepackaged PDMS MFCs are 1.7 cm × 1.7 cm × 0.2 cm. Thevolume of the electrode compartment is 15-μL. The electrode



Fig. 6. Assembly and packaging of PDMS MFC.

surface-area-to-volume ratio is 144 cm−1 and is calculatedby dividing the effective electrode area by the compartmentvolume. In comparison to previous silicon microchannel MFCwith a 0.51-cm2 effective electrode area and a 16-μL volume[58], the PDMS MFC has 4.5 times increase in the surface-area-to-volume ratio.

IV. EXPERIMENTAL SETUP

S. cerevisiae (Sigma YSC1) is nonpathogenic and easy toculture [87]. S. cerevisiae of 85 g/L dehydrated weight perbuffer volume was autolyzed in a 0.1-M potassium phosphatebuffer (pH 7.0) at 37 ± 0.5 ◦C for 12 h. The potassium phos-phate buffer was prepared from American Chemical Society(ACS) grade potassium phosphate diabasic anhydrous (FisherScientific, BP363) and potassium phosphate monobasic (FisherScientific, P275) dissolved in DI water with 18.2-MΩ · cmresistivity. Before the experiment, the microbial culture waswashed twice in phosphate buffer. Then, it was mixed with10-mM MB (Fisher Scientific, M291, certified for use inbacteriology and bloodstain) in pH 7.0 phosphate buffer tobecome the phosphate-buffered anolyte. Finally, 1-M glucose(D+ glucose, Acros Organics, AC41095, ACS) was addedto the medium. The catholyte was prepared using the samepH 7.0 phosphate buffer, composed of 20-mM potassiumferri(III)cyanide (Acros Organics, AC424125000, ACS).

The human plasma for the power generation experimentwas prepared using the following procedures. Human plasma(Sigma, P9532) was used both as the electrolyte and as the fuelsupply. The acidity of the human plasma was measured to bepH 7.4 (Fisher Scientific, Accumet AB15), and the concentra-tions of blood glucose, protein, nitrite, and leukocytes wereshown to be in the normal physiological range (DiaScreen4NL, hypoguard) [28]. The plasma glucose concentration was75 mg/L (4.2 mM), measured by colorimetric analysis [88].Dehydrated S. cerevisiae was cultured in human plasma in an85-g/L dry weight ratio at 37 ± 0.5 ◦C for 12 h and washedtwice with fresh plasma. The harvested S. cerevisiae was thenresuspended in 10-mM MB dissolved in human plasma asthe anolyte. The catholyte was prepared by dissolving 20-mMFerri(III)cyanide in the human plasma.

The experiments were conducted at 37 ± 0.5 ◦C inside an in-cubation chamber to simulate in vivo human body temperature.

Fig. 7. SEM micrograph showing the S. cerevisiae cultured on the PDMSelectrode after 18 h. The S. cerevisiae (mother cell) is surrounded by acluster of newborn cells. The mother cell with buds on the surface shows thereproductive characteristic of ascomycetous microorganism. The ultrastructuresof the developing newborn cell shrink due to dehydration inside the low-pressure SEM chamber.

The temperature of the MFC was monitored by a K-type ther-mal couple (Circuit Test Electronic, TL180) directly adheredon top of the external surface of the anodic compartment andconnected to the microprocessor thermometer (HH23, Omega)with ±0.1 ◦C accuracy. The PDMS MFC is preheated to 37 ◦Cinside an incubation chamber for 1 h before the experiment.Electrolytes were injected by micropipette to the anode andcathode compartments. Electrical potential and current outputswere measured using a data acquisition system (NI 4070 andLab View 7.1, National Instruments) at a 2-Hz sampling rate.The voltage and current resolution were 1 μV and 10 nA,respectively. Four different resistors, 47 Ω, 100 Ω, 470 Ω,and 1 kΩ (Velleman, K/RES-E3, 5% tolerance) were used inboth the phosphate buffer and human plasma as the dischargeloads. To ensure the consistency of the electrolyte and preventresidues from remaining inside the fuel cell compartments,fresh cultured medium and a new PDMS MFC were used foreach test.

V. PERFORMANCE OF THE PDMS MFC

A. Microbial Adaption

The morphology of the colonized solid surface significantlyaffects the viability of S. cerevisiae [61]–[63]. Qualitativeanalysis of the growth of S. cerevisiae on the gold-evaporatedPDMS electrode was performed using a SEM. Five piecesof samples were cultured with 85-g/L dry weight ratioS. cerevisiae in pH 7.0 phosphate buffer for 18 h at 37 ± 0.5 ◦C.All samples were observed to have their entire surface colo-nized by S. cerevisiae. When S. cerevisiae grows to a criticalcell volume about 30–50 μm3, it initiates buds for reproduction[89]. S. cerevisiae adopts asymmetric cell division for buddingwith the newborn cells being smaller than the mother cell[90], [91, pp. 101–183]. Fig. 7 shows a SEM picture of anS. cerevisiae mother cell surrounded by a cluster of newborncells on the PDMS electrode. The mother cell is undergoingreproductive cell division with buds extruded from its surface.



Fig. 8. OCP of the MFC under different anolytes. The catholytes are 20-mMpotassium ferricyanide in pH 7.0 phosphate buffer.

B. Effect of Anolyte Constituents on OCP

Microbial metabolic processes are significantly influenced byglucose concentration, condition of the culture medium, andamount of S. cerevisiae inside the anolyte [74], [91, pp. 52–61].These effects are reflected in the OCP response [36], [46], [92].Fig. 8 shows the OCP obtained by different anolyte combina-tions in pH 7.0 phosphate buffer. The MFC with 10-mM MB,S. cerevisiae, and 1-M glucose as the anolyte had the high-est OCP throughout the experiment. Initially, the OCP was475.3 mV. As the fuel was consumed, the OCP graduallydecreased to 400.9 mV after 60 min, with an average OCPof 421.8 mV. The MFC without MB had an average OCP of107.3 mV, which is 25.4% of the OCP with MB. As the metabo-lized electrons were barely liberated from the microbial interiorwithout MB, the potential induced on the anode may come fromthe interface between the microbial membrane surface and theelectrode. The cell membrane of the microorganism generates−120 to −160 mV as the driving force to release protons[78]. The OCP from the anolyte with only phosphate bufferand 1-M glucose showed an average background potential of7.1 μV, which may be attributed to the oxidization of glucoseto gluconic acid catalyzed on the gold electrode [93]

C6H12O6 + OH− Au−−−→ C6H12O7 + H+ + 2e−. (9)

Other studies have shown that when the anolyte contains only10-mM MB in a pH 7.0 phosphate buffer without S. cerevisiae,the OCP is sustained at around 40 mV [58].

Theoretically, the OCP of the MFC can be evaluated from theoverall redox reaction by the Nernst equation [33], [86], [94]

OCP = OCP0 − RT

nFIn

Π(activities of reductant)Π(activities of oxident)

(10)

where R = 8.314 J · mol−1 · K−1 is the ideal gas constant,T = 310.15 K (37 ◦C) is the operating temperature measuredin kelvins, n = 4 is the number of moles of electron involvedin (8), F = 96 485.3 C/mol is the Faraday constant, and OCP0

is the OCP at standard equilibrium condition, which is calcu-lated as

OCP0 = E0cathode − E0

anode = 0.965 V. (11)

From (10),

OCP = 0.965 − RT

nFIn

[MB]2[H20]2

[MBH]2[O2][H+]2. (12)

From (3), ([MB]/[MBH]) is equal to ([NAD+]/[NADH]);therefore, (12) can be modified to

OCP = 0.965 − RT

nFIn

[NAD+]2[H20]2

[NADH]2[O2][H+]2(13)

where the activity of water in liquid is 1, the activity of oxygenis the ratio of partial pressure over standard atmospheric pres-sure, and the activity ratio of NAD+ and NADH, which is equalto the intracellular concentration ratio of NAD+ and NADH,is equal to 1.48 [86], [94], [95]. The relationship between pHvalue and proton activity is found as [94]

In1

[H+]= 2.303 log10

1[H+]

= 2.303 pH. (14)

The Nernst equation then becomes

OCP = 0.965 − 2.303RT

2FpH − RT

2FIn

1.48[PO2]

. (15)

With the assumption that the partial pressure of oxygen PO2

is equivalent to 0.0619 atm at 37 ◦C in water [96], [97], thetheoretical OCP is 0.71 V. The difference of the measured OCPfrom the theoretical value may be attributed to the irreversibleloss on the electrode interfaces, ohmic resistances, concentra-tion gradients, the redox reversibility of the mediator, or thephysiology of the microorganism [21], [32], [34], [36], [40],[48]. One of the factors affecting the decline of OCP is thechange of the redox potential of MB as the medium becomesacidic. An experiment showed the increase redox potential ofMB from 0.011 to 0.101 V when the acidity of the anolytechanges from pH 7.0 to pH 5.0 at 30 ◦C [48]. This reducesthe OCP as in (11). The alteration of the medium acidity comesfrom the accumulation of carbon dioxide in our MFC design(1), regulated release of protons from the microbia [78], andthe gold-catalyzed gluconic acid [93].

Fig. 9 shows the OCP as a function of glucose concentration.The data refer to average OCP values in a 60-min operation inpH 7.0 phosphate buffer with 10-mM MB. The curve showsa rapid increase from 2.5-mM to 0.25-M glucose and be-comes steady at 1-M concentration. A regression analysis withR-square value of 0.98 showed a correlation between the OCPand the logarithmic glucose concentration. Other MFC studieshave also reported different degrees of OCP variation with gluc-ose or other substrate concentrations [21], [27], [36], [98], [99].The Monod growth kinetic model gives a general relation between microbial growth and the substrate concentration as [100]

μ = μmax

(s

Ks + s

)(16)



Fig. 9. Influence of glucose concentration on OCP. The experimental data arecompared with modified Monod growth kinetic model. The inset semilog plotshows a correlation between OCP and logarithmic glucose concentration.

Fig. 10. Current density generated from a single drop of 15-μL anolyte inpH 7.0 phosphate buffer at different loads. The anolyte contains 1-M glucoseand 10-mM MB with S. cerevisiae. The catholyte contains 20-mM potassiumferrocyanide in 15-μL 0.1-M phosphate buffer.

where μ is the microbial specific growth rate, μmax is themaximum specific growth rate, s is the substrate concentration,and Ks is the substrate affinity constant. A modification ofthe Monod growth kinetic model correlated to OCP has beenproposed for MFC with the following equation [99]:

OCP = OCPmax

(s

Ks + s

)(17)

where OCPmax is the maximum voltage measured at the highestglucose concentration performed in the experiment and Ks

is determined empirically. The simulation curve, using (17),is shown in Fig. 9 and compared to the measured OCP. Theempirical value of Ks is 3.0 g/L, and the R-square value is 0.95,from the curve fitting.

C. Current and Power Densities

Fig. 10 shows the current density from the four differentloads using phosphate as the electrolyte. The current densityJ is defined as

J =I

A(18)

Fig. 11. Characteristics of MFC average power density powered by a singledrop of 15-μL anolyte with 1-M glucose solution.

where I is the instantaneous current and A is the geometric areaof the electrodes (1.2 cm2) [21]. Each current density shows anincrease at the beginning and then a steep decline. This profilecan be described by the chemical balance between reactions (5)and (6). Before discharging, the cathode compartment containsonly Fe(CN)3

−6 ions. When the electrons start to flow from the

anode to the cathode via the external circuit, the Fe(CN)3−

6 ionsare being reduced to Fe(CN)4

−6 ions, and the current density

increases. Because of the inefficient proton reduction fromFe(CN)4

−6 ions; however, more Fe(CN)4

−6 ions accumulate in

the cathode compartment and fewer Fe(CN)3−

6 ions remainto react with the electrons [46], [87]. As a result, the currentdensity decreases. Another factor accounting for the decline isthe accumulation of protons that changes the pH value of thesystem and denatures the electron mediator [23], [40], [48].This decline may also be due to the limited fuel volume sothat electron-discharged rate is much higher than the generationrate. It may also be related to the microbial growth rate in theanolyte during the discharge process. A specific analysis wouldbe required for better understanding.

The highest current density measured was 34.0 μA/cm2

(100 Ω), and the lowest was 17.4 μA/cm2 (1 kΩ). The averagecurrent densities for 60 min were 8.2, 5.1, 2.7, and 1.6 μA/cm2

for 47 Ω, 100 Ω, 470 Ω, and 1 kΩ loads, respectively. A controlexperiment with 100 Ω shows the background current densitywhen the MFC is operated without S. cerevisiae and MB in theanolyte.

Fig. 11 shows the average power density with different loads.The average power density Pave is calculated by

Pave =R

tA

t∫0

I2dt (19)

where R is the load resistance and t is the discharge duration.The average power density decreased with time as the currentdensity decreased. The results show the highest measuredpower density output at around 500 Ω. According to the MFCimpedance model, the output power of the MFC is maximizedwhen the MFC internal resistance is equal to the external load



TABLE ICOMPARISON OF MICROFABRICATED MFC CHARACTERISTICS IN

PHOSPHATE BUFFER MEDIUM

Fig. 12. OCP development of the MFCs operating with human blood plasmaas the glucose supply and electrolyte in the anode compartment.

[101]. This implies that the PDMS MFC in this phosphatebuffer condition has an internal resistance at around 500 Ω.Table I summarizes the performance of the PDMS MFC andcompares it with a silicon-based micromachined MFC with azigzag microchannel design [58]. Both MFCs were catalyzedby S. cerevisiae. The compositions of the anolyte and catholytein both MFCs were the same, but the silicon micromachinedMFC has four times the culture time in a full nutrient conditionand connected with a 101-Ω load [58]. During the 40-min dis-charge period, the PDMS MFC had an average power densityof 8.1 nW/cm2 at 100-Ω load, which is 40.5 times greater thanthat of the silicon micromachined MFC.

D. Power From Human Plasma

Fig. 12 shows the OCP of a PDMS MFC using human plasmaas the electrolyte. The MFC with S. cerevisiae and MB hadan initial maximum OCP of 488.1 mV and a minimum of258.6 mV after 60 min. The average OCP was 297.4 mV. Thedecline in OCP occurred in the first 5 min and then steadily de-creased. The initial decline may account for protein absorptionon the proton exchange membrane prohibiting proton trans-mission [102], [103]. Two control experiments using different

anolyte compositions were conducted: 1) without S. cerevisiaeand 2) without both S. cerevisiae and MB. The average OCPswere 198.0 and 198.4 mV, respectively.

Fig. 13 shows the current density of PDMS MFC usinghuman plasma as the electrolyte. The advantage of usinghuman plasma rather than phosphate as the electrolyte isthat the current density drives from both the biocatalyticeffect of the blood glucose and the secondary redox reactionfrom the ionic constituents inside the human plasma. Thissecondary reaction includes some amino acid oxidations on theanode with negative redox potentials such as [GSH]/[GSSG](Eo = −240 mV at pH 7.0), [mono(des-Glu)-GSSG]/[GSH-][CysGly] (Eo = −275 mV at pH 7.4), [CySSG]/[GSH][Cys](Eo = −275 mV at pH 7.4), and [CySS]/[Cys]2 (Eo =−250 mV at pH 7.4), where GSH is a cysteine-containingtripeptide glutathione, GSSG is glutathione disulfide, CysGly isL-cysteinylglycine, Cys is cysteine, CySS is cystine, and Eo isthe standard redox potential [104], [105]. The current densitieswithout both S. cerevisiae and MB in human plasma are alsoshown in Fig. 13. The 60-min average current densities fromdischarge loads of 47 Ω, 100 Ω, 470 Ω, and 1 kΩ were 8.8, 8.0,6.7, and 4.3 μA/cm2, respectively. A 56%–71% of the currentdensities were contributed by the biocatalytic effect of bloodglucose.

Fig. 14 shows the average power density as a function of dis-charge time and loads. The highest average power density out-puts were 140.1 and 42.4 nW/cm2 for 1 kΩ at 15- and 60-mindurations, respectively. Compared to Fig. 11, the average powerdensity in human plasma increases from 47 Ω to 1 kΩ, whichimplies that the internal resistance of the MFC may be over1 kΩ in the human plasma culturing condition [101]. Severalstudies suggest that the anodic resistance dominates the internalresistance [24], [106] and further analysis from electrochemicalimpedance spectroscopy would provide a better understanding[101]. Table II compares the electrical performances of theMFC operating in a phosphate buffer versus human plasma asthe electrolyte. Although the average OCP from human plasmawas lower than that from the phosphate buffer electrolyte,human plasma generated a higher average current density andpower density. As a possible explanation, human plasma con-tains comprehensive minimal salts which prompts the growthand metabolism of S. cerevisiae. For example, the existence ofmagnesium ion prompts the synthesis of transphosphorylationenzymes, and potassium ion can activate the metabolism ofpyruvate in glucose conversion [91, pp. 52–61], [107].

E. Coulombic Efficiency of MFC

Coulombic efficiency is a measure of biocatalytic electronconversion from the fuel to electricity [22], [34], [40], [41].Theoretically, complete oxidation of a glucose molecule ex-tracts 24 electrons as [34], [58]

C6H12O6 + 6H20 → 6CO2 + 24H+ + 24e−. (20)

Therefore, the maximum electron yield Y0, with the volumeof anode compartment L and the glucose concentration Cg , is



Fig. 13. Current density performance of PDMS MFC powered by 15-μL human blood plasma in the anode compartment for 60-min operation under variousdischarge loads: (a) 47 Ω, (b) 100 Ω, (c) 470 Ω, and (d) 1 kΩ. The electrolytes are identical in each case, with anolyte: 15-μL human plasma and 10-mM MBwith S. cerevisiae; and catholyte: 20-mM potassium ferrocyanide in 15-μL human plasma.

Fig. 14. Average power density produced from the glucose content of 15-μLhuman blood plasma with a normal blood glucose concentration. The legend“gross” shows the total power density measured from the MFC, and “net” refersto the contribution solely from the blood glucose biocatalyzed by S. cerevisiae.

expressed as [58]

Y0 = 24 × 6.023 × 1023 × 1.6 × 10−19LCg (21)

where 6.023 × 1023 is the Avogadro number of electrons permole and 1.6 × 10−19 is the coulomb of charge of a single

electron. Y0’s for the phosphate buffer and human plasmaexperiments are calculated as 34.7 and 0.146 C, respectively.Practically, electron liberation is restricted by the biocatalyticrate, shuttling efficiency of the electron mediator, and the me-diator redox rate on the electrode [21], [34], [40]. In addition,some electrons are used by the microorganism to maintain thevitality through ATP synthesis [34]. The coulombs of electronsextracted for electricity supply Y is obtained by integrating thecurrent produced from biocatalytic effect with respect to time.The coulombic efficiency CE is defined as the percentage ratiobetween coulombs of electron flow through the external circuitto the maximum electron yield as follows [40]:

CE =

∫ t

0 Idt

Y0× 100%. (22)

Table II summarizes the coulombic efficiency of the PDMSMFCs operating for 60 min. The coulombic efficiency derivedfrom human plasma ranges from 9.0% to 14.7%, which ismuch higher than that of the phosphate buffer condition with0.02%–0.10%. Compared with other macroscale MFCs, withreported 50% to roughly 100% coulombic efficiency, theiroperating durations of MFC are unrestricted, varying from hourto days of operation [22], [34], [55].



TABLE IICOMPARISON OF PDMS MFC CHARACTERISTICS IN PHOSPHATE BUFFER VERSUS HUMAN PLASMA MEDIUM

VI. CONCLUSION

This paper has presented a microfabricated PDMS MFC thatuses an embedded micropillar structure to increase electricaloutput. The structure and packaging material of the MFC con-sist entirely of PDMS making the fuel cell potentially suitablefor body implantation. The MFC has a maximum OCP of488.1 mV, current density of 30.2 μA/cm2, and power densityof 401.2 nW/cm2. With a testing time of 60 min, the MFCproduces an average OCP of 297.4 mV, average current densityof 4.3 μA/cm2, and an average power density of 42.4 nW/cm2

under 1-kΩ load. The coulombic efficiency for electron con-version in 60 min ranges from 9% to 14.7%. In comparisonwith the silicon micromachined MFCs, the PDMS MFC has a4.5 times increase in electrode surface-area-to-volume ratio and40.5 times increase in power density.

Experimental studies of MFC generating power from a singledroplet of 15-μL human plasma partially demonstrate the pos-sibility of in vivo electricity generation. The demonstration of aflexible PDMS MFC is one step further toward the ultimate goalof a long-term miniature power source for implantable devices.

For practical applications, the future implantable MFC shouldbe able to exchange blood and carbon dioxide between thefuel cell interior and exterior, and prevent leaching of electronmediator, oxidant, and microbia to the blood stream. Steadystate performance of the in vivo power generation should beanalyzed. Design optimization of the micropillar quantity andgeometry may further increase electron harvesting. Analysis ofthe protein absorption mechanism, modification of the protonexchange membrane may also increase the power output ofMFC. In addition, other investigations in biocatalyst reactionand diffusion rate, electron mediator interaction, electrode im-pedances, and oxidant reaction response can further advancethe coulombic efficiency and longevity of this new-generationpower source.

ACKNOWLEDGMENT

The PDMS MFC presented in this paper was fabricated inthe Advanced Materials and Process Engineering Laboratory(AMPEL) and the MEMS & Nano Engineering Laboratory.



REFERENCES

[1] A. C. R. Grayson, R. S. Shawgo, A. M. Johnson, N. T. Flynn, Y. Li,M. J. Cima, and R. Langer, “A BioMEMS review: MEMS technology forphysiologically integrated devices,” Proc. IEEE, vol. 92, no. 1, pp. 6–21,Jan. 2004.

[2] P. S. Motta and J. W. Judy, “Multielectrode microprobes for deep-brainstimulation fabricated with a customizable 3-D electroplating process,”IEEE Trans. Biomed. Eng., vol. 52, no. 5, pp. 923–933, May 2005.

[3] R. Birana, D. C. Martin, and P. A. Tresco, “Neuronal cell loss ac-companies the brain tissue response to chronically implanted siliconmicroelectrode arrays,” Exp. Neurol., vol. 195, no. 1, pp. 115–126,Sep. 2005.

[4] W. Mokwa, “Medical implants based on microsystems,” Meas. Sci. Tech-nol., vol. 18, no. 5, pp. R47–R57, Mar. 2007.

[5] J. U. Meyer, “Retina implant—A bioMEMS challenge,” Sens. ActuatorsA, Phys., vol. 97/98, pp. 1–9, Apr. 2002.

[6] W. Liu, K. Vichienchom, M. Clements, S. C. DeMarco, C. Hughes,E. McGucken, M. S. Humayun, E. De Juan, J. D. Weiland, andR. Greenberg, “A neuro-stimulus chip with telemetry unit for reti-nal prosthetic device,” IEEE J. Solid-State Circuits, vol. 35, no. 10,pp. 1487–1497, Oct. 2000.

[7] J. F. Rizzo, III, J. Wyatt, J. Loewenstein, S. Kelly, and D. Sbire, “Percep-tual efficacy of electrical stimulation of human retina with a microelec-trode array during short-term surgical trials,” Investig. Ophthalmol. Vis.Sci., vol. 44, no. 12, pp. 5362–5369, Dec. 2003.

[8] D. D. Zhou and R. J. Greenberg, “Microsensors and microbiosensors forretinal implants,” Front Biosci., vol. 10, pp. 166–179, Jan. 2005.

[9] D. C. Rodger, A. J. Fong, W. Li, H. Ameri, I. Lavrov, H. Zhong,S. Saati, P. Menon, E. Meng, J. W. Burdick, R. R. Roy, V. R. Edgerton,J. D. Weiland, M. S. Humayun, and Y. C. Tai, “High-density flexibleparylene-based multielectrode arrays for retinal and spinal cord stimu-lation,” in Proc. IEEE Int. Conf. Solid-State Sens., Actuators Microsyst.Transducers, Lyon, France, 2007, pp. 1385–1388.

[10] T. E. Bell, K. D. Wise, and D. J. Anderson, “A flexible micromachinedelectrode array for a cochlear prosthesis,” Sens. Actuators A, Phys.,vol. 66, no. 1–3, pp. 63–69, Apr. 1998.

[11] J. Wang, M. Gulari, and K. D. Wise, “An integrated position-sensingsystem for a MEMS-based cochlear implant,” in IEDM Tech. Dig., 2005,pp. 129–132.

[12] M. A. Zurcher, D. J. Young, M. Semaan, C. A. Megerian, andW. H. Ko, “MEMS middle ear acoustic sensor for fully implantablecochlear prosthesis,” in Proc. 20th IEEE MEMS Conf., 2007, pp. 11–14.

[13] J. U. Meyer, M. Schuttler, H. Thielecke, and T. Stieglitz, “Biomedicalmicrodevices for neural interfaces,” in Proc. IEEE Microtechnol. Med.Biol., 2000, pp. 447–453.

[14] K. C. Cheung and P. Renaud, “BioMEMS for medicine: On-chip cellcharacterization and implantable microelectrodes,” Solid State Electron.,vol. 50, no. 4, pp. 551–557, Apr. 2006.

[15] D. Purves, G. J. Augustine, D. Fitzpatrick, L. C. Katz, A. S. LaMantia,J. O. McNamara, and S. M. Williams, Eds., “Electrical signal of nervecell and voltage-dependent membrane permeability,” in Neuroscience,2nd ed. Sunderland, MA: Sinauer Associates Inc. 2001, ch. 2–3,pp. 31–68.

[16] V. S. Mallela, V. Ilankumaran, and N. S. Rao, “Trends in cardiac pace-maker batteries,” Indian Pacing Electrophysiol. J., vol. 4, no. 4, pp. 201–212, Oct. 2004.

[17] C. K. Dyer, “Fuel cells and portable electronics,” in VLSI Symp. Tech.Dig., 2004, pp. 124–128.

[18] A. Heller, “Potentially implantable miniature batteries,” Anal. Bioanal.Chem., vol. 385, no. 3, pp. 469–473, Jun. 2006.

[19] P. B. Koeneman, I. J. Busch-Vishniac, and K. L. Wood, “Feasibility ofmicro power supplies for MEMS,” J. Microelectromech. Syst., vol. 6,no. 4, pp. 355–362, Dec. 1997.

[20] S. B. Dunkerton and S. Hodgins, “Healthy aims-development of im-plantable microsystems medical devices,” in Proc. ASM Int. Mater.Process. Med. Devices Conf., St. Paul, MN, 2004, pp. 161–165.

[21] E. Katz, A. N. Shipway, and I. Willner, “Biochemical fuel cell,” inHandbook of Fuel Cells, vol. 1, W. Vielstich, H. A. Gasteiger, andA. Lamm, Eds. Hoboken, NJ: Wiley, 2003, ch. 21, pp. 1–28.

[22] K. Rabaey, G. Lissens, and W. Verstraete, “Microbial fuel cells: Perfor-mances and perspectives,” in Biofuels for Fuel Cells: Biomass Fermen-tation Towards Usage in Fuel Cell, P. Westermann, P. Haberbauer, andM. Lens, Eds. London, U.K.: IWA Publ., 2005, ch. 20. sec 20.7.1.

[23] G. A. Justin, Y. Zhang, M. Sun, and R. Sclabassi, “Biofuel cells: Apossible power source for implantable electronic devices,” in Proc. 26thAnnu. Int. Conf. IEEE EMBS, San Francisco, CA, 2004, pp. 4096–4099.

[24] M. Grzebyk and G. PoŸniak, “Microbial Fuel Cells (MFCs) with in-terpolymer cation exchange membranes,” Sep. Purif. Technol., vol. 41,no. 3, pp. 321–328, Feb. 2005.

[25] A. F. Ghenciu, “Review of fuel processing catalysts for hydrogen pro-duction in PEM fuel cell systems,” Curr. Opin. Solid State Mater. Sci.,vol. 6, no. 5, pp. 389–399, Oct. 2002.

[26] J. Larminie and A. Dicks, Fuel Cell Systems Explained, 2nd ed.Chichester, U.K.: Wiley, 2003, pp. 238–307.

[27] A. L. Walker and C. W. Walker, “Biological fuel cell and an applicationas a reserve power source,” J. Power Sources, vol. 160, no. 1, pp. 123–129, Sep. 2006.

[28] J. M. Orten, Human Biochemistry, 10th ed. St. Louis, MO: Mosby,1982, pp. 631–632.

[29] J. Bolincier, U. Ungerstedt, and P. Arner, “Microdialysis measurementof the absolute glucose concentration in subcutaneous adipose tissueallowing glucose monitoring in diabetic patients,” Diabetologia, vol. 35,no. 12, pp. 1177–1180, Dec. 2004.

[30] S. Kerzenmacher, R. Sumbharaju, J. Ducree, R. Zengerle, andF. von Stetten, “A surface mountable glucose fuel cell for medical im-plants,” in Proc. IEEE Int. Conf. Solid-State Sens., Actuators Microsyst.Transducers, Lyon, France, 2007, pp. 125–127.

[31] P. R. Donald, C. Malan, and A. van der Walt, “Simultaneous determi-nation of cerebrospinal fluid glucose and blood glucose concentrationsin the diagnosis of bacterial meningitis,” J. Pediatr., vol. 103, no. 3,pp. 413–415, Sep. 1983.

[32] L. B. Wingard, C. H. Shaw, and J. F. Castner, “Bioelectrochemi-cal fuel cells,” Enzyme Microb. Technol., vol. 4, no. 3, pp. 137–142,May 1982.

[33] R. A. Bullen, T. C. Arnot, J. B. Lakeman, and F. C. Walsh, “Biofuel cellsand their development,” Biosens. Bioelectron., vol. 21, no. 11, pp. 2015–2045, May 2006.

[34] U. Schröder, “Anodic electron transfer mechanisms in microbial fuelcells and their energy efficiency,” Phys. Chem. Chem. Phys., vol. 9,no. 21, pp. 2619–2629, Jun. 2007.

[35] D. R. Lovley, “Bug juice: Harvesting electricity with microorganisms,”Nat. Rev. Microbiol., vol. 4, no. 7, pp. 497–508, Jul. 2006.

[36] M. C. Potter, “Electrical effects accompanying the decomposition of or-ganic compounds,” Proc. R. Soc. Lond., B Contain. pap. biol. Character,vol. 84, no. 571, pp. 260–286, Sep. 1911.

[37] B. Cohen, “The bacterial culture as an electrical half-cell,” J. Bacteriol.,vol. 21, no. 1, pp. 18–19, 1931.

[38] J. B. Davis, “Generation of electricity by microbial action,” Adv. Appl.Microbiol., vol. 5, pp. 51–64, 1963.

[39] M. J. Allen, “Biofuel cells,” in Methods in Microbiology, J. R. Norrisand D. W. Ribbon, Eds. New York: Academic, 1972, pp. 247–283.

[40] J. L. Stirling, H. P. Bennetto, G. M. Delaney, J. R. Mason, S. D. Roller,K. Tanaka, and C. F. Thurston, “Microbial fuel cells,” Biochem. Soc.Trans., vol. 11, no. 4, pp. 451–453, Aug. 1983.

[41] H. P. Bennetto and J. L. Stirling, “Anodic reactions in microbial fuelcells,” Biotechnol. Bioeng., vol. 25, no. 2, pp. 559–568, Feb. 1983.

[42] K. Tanaka, C. A. Vega, and R. Tamamushi, “Mediating effects of ferricchelate compounds in microbial fuel cells,” Bioelectrochem. Bioenerg.,vol. 11, pp. 135–143, 1983.

[43] S. D. Roller, H. P. Bennetto, G. M. Delaney, J. R. Mason, J. L. Stirling,and C. F. Thurston, “Electron-transfer coupling in microbial fuel cells:1. Comparison of redox-mediator reduction rates and respiratory ratesof bacteria,” J. Chem. Technol. Biotechnol., vol. 34, no. 1, pp. 3–12,1984.

[44] R. M. Allen and H. P. Bennetto, “Microbial fuel cells: Electricity pro-duction from carbohydrates,” Appl. Biochem. Biotechnol., vol. 39/40,pp. 27–40, 1993.

[45] V. G. Debabov, “Electricity from microorganisms,” Microbiology,vol. 77, no. 2, pp. 123–131, Apr. 2008.

[46] G. M. Delaney, H. P. Bennetto, J. R. Mason, S. D. Roller, J. L. Stirling,and C. F. Thurston, “Electron-transfer coupling in microbial fuel cells. 2.Performance of fuel cells containing selected microorganism-mediator-substrate combinations,” J. Chem. Technol. Biotechnol., vol. 34B,pp. 13–27, 1984.

[47] G. M. Walker, Yeast Physiology and Biotechnology. New York: Wiley,1998, pp. 22–23.

[48] L. F. Hewitt, Oxidation-Reduction Potentials in Bacteriology and Bio-chemistry. London, U.K.: County Council, 1933, pp. 1–81.

[49] G. Daemen, R. G. H. Catharina, and P. H. P. Groeneveld, “Methyleneblue increases myocardial function in septic shock,” Crit. Care. Med.,vol. 23, no. 8, pp. 1363–1370, Aug. 1995.

[50] L. Aurelian, “The ‘viruses of love’ and cancer,” Amer. J. Med. Technol.,vol. 40, no. 11, pp. 496–512, Nov. 1974.



[51] J. F. Papina, R. A. Floydb, and D. P. Dittmerc, “Methylene blue photoin-activation abolishes West Nile virus infectivity in vivo,” Antivir. Res.,vol. 68, no. 2, pp. 84–87, Nov. 2005.

[52] D. M. Roberts and N. A. Buckley, “Pharmacokinetic considerationsin clinical toxicology: Clinical applications,” Clin. Pharmacokinet.,vol. 46, no. 11, pp. 897–939, 2007.

[53] H. Nishioka, “Mutagenic activities of metal compounds in bacteria,”Mutat. Res., vol. 31, no. 3, pp. 185–189, Jun. 1975.

[54] P. Hantson, P. N. Geye, M. Laforge, J. L. Clemessy, and F. Baud,“Suicide attempt by ingestion of potassium ferricyanide,” Clin. Toxicol.,vol. 34, no. 4, pp. 471–473, 1996.

[55] S. K. Chaudhuri and D. R. Lovely, “Electricity generation by direct oxi-dation of glucose in mediatorless microbial fuel cells,” Nat. Biotechnol.,vol. 21, no. 10, pp. 1229–1232, 2003.

[56] H. J. Kim, H. S. Park, M. S. Hyum, I. S. Chang, M. A. Kim, andB. H. Kim, “A mediatorless microbial fuel cell using a metal reduc-ing bacterium, Shewanella putrefacians,” Enzyme Microbiol. Technol.,vol. 30, pp. 145–152, 2002.

[57] C. R. Myers and M. J. Myers, “Localization of cytochromes to the outermembrane of anearobically grown Shewanella putrefaciens MR-1,”J. Bacteriol., vol. 174, no. 11, pp. 3429–3438, 1992.

[58] M. Chiao, K. B. Lam, and L. Lin, “Micromachined microbial and photo-synthetic fuel cells,” J. Micromech. Microeng., vol. 16, no. 12, pp. 2547–2553, Oct. 2006.

[59] K. B. Lam, E. A. Johnson, M. Chiao, and L. Lin, “A MEMS photosyn-thetic electrochemical cell powered by subcellular plant photosystems,”J. Microelectromech. Syst., vol. 15, no. 5, pp. 1243–1250, Oct. 2006.

[60] Z. Nie and E. Kumacheva, “Patterning surfaces with functional poly-mers,” Nat. Mater., vol. 7, no. 4, pp. 277–290, 2008.

[61] E. O. Morris and J. S. Hough, “Some aspects of the giant colony charac-teristics of Saccharomyces cerevisiae,” J. Inst. Brew., vol. 62, pp. 466–469, 1956.

[62] C. J. Gimeno, P. O. Ljungdahl, C. A. Styles, and G. R. Fink, “Unipolarcell divisions in the yeast S. cerevisiae lead to filamentous growth:Regulation by starvation and RAS,” Cell, vol. 68, no. 6, pp. 1077–1090,Mar. 1992.

[63] R. S. Kamath and H. R. Bungay, “Growth of yeast colonies on solidmedia,” J. Gen. Microbiol., vol. 134, no. 11, pp. 1411–1416, Nov. 1988.

[64] H. Kudoa, T. Sawadaa, E. Kazawab, H. Yoshidab, Y. Iwasakic, andK. Mitsubayashia, “A flexible and wearable glucose sensor based onfunctional polymers with Soft-MEMS techniques,” Biosens. Bioelec-tron., vol. 22, no. 4, pp. 558–562, Oct. 2006.

[65] K. J. Wynne and J. M. Lambert, “Silicones,” in Encyclopedia of Bioma-terials and Biomedical Engineering, G. L. Bowlin and G. Wnek, Eds.New York: Marcel Dekker, 2004, pp. 1348–1362.

[66] S. Hu, X. Ren, M. Bachman, C. E. Sims, G. P. Li, and N. L. Allbritton,“Surface modification of poly(dimethylsiloxane) microfluidic devices byultraviolet polymer grafting,” Anal. Chem., vol. 74, no. 16, pp. 4117–4123, Jun. 2002.

[67] S. K. Sia and G. M. Whitesides, “Microfluidic devices fabricated inPoly(dimethylsiloxane) for biological studies,” Electrophoresis, vol. 24,no. 21, pp. 3563–3576, Nov. 2003.

[68] A. C. R. Grayson, R. S. Shawgo, Y. Li, and M. J. Cima, “ElectronicMEMS for triggered delivery,” Adv. Drug Deliv. Rev., vol. 56, no. 2,pp. 173–184, Feb. 2004.

[69] M. C. Bélanger and Y. Marois, “Hemocompatibility, biocompatibility,inflammatory and in vivo studies of primary reference materials low-density polyethylene and polydimethylsiloxane: A review,” J. Biomed.Mater. Res., vol. 58, no. 5, pp. 467–477, Jul. 2001.

[70] M. T. Khorasani and H. Mirzadeh, “In vitro blood compatibility ofmodified PDMS surfaces as superhydrophobic and superhydrophilic ma-terials,” J. Appl. Polym. Sci., vol. 91, no. 3, pp. 2042–2047, 2004.

[71] K. Shah, W. C. Shin, and R. S. Besser, “A PDMS micro proton exchangemembrane fuel cell by conventional and non-conventional microfabrica-tion techniques,” Sens. Actuator B, Chem., vol. 97, no. 2/3, pp. 157–167,Feb. 2004.

[72] E. R. Choban, L. J. Markoski, A. Wieckowski, and P. J. A. Kenis, “Mi-crofluidic fuel cell based on laminar flow,” J. Power Sources, vol. 127,no. 1, pp. 54–60, Mar. 2004.

[73] Y. Jiang, X. Wang, K. Xie, X. Qiu, L. Zhong, and L. Liu, “A micro directmethanol fuel cell using PDMS assembly technology,” in Proc. IEEE Int.Conf. Solid-State Sens., Actuators Microsyst. Transducers, Seoul, Korea,2005, pp. 303–306.

[74] J. A. Barnett, “The utilization of sugars by yeasts,” Adv. Carbohydr.Chem. Biochem., vol. 32, pp. 125–234, 1976.

[75] M. J. Pelczar, E. C. S. Chan, and N. R. Krieg, Microbiology, 5th ed.New York: McGraw-Hill, 1986, pp. 171–195.

[76] K. Rabaey and W. Verstraete, “Microbial fuel cells: Novel biotechnologyfor energy generation,” Trends Biotechnol., vol. 23, no. 6, pp. 291–298,Jun. 2005.

[77] M. J. B. Hauser, A. Lunding, and L. F. Olsen, “On the role of methyleneblue in the oscillating peroxidase-oxidase reaction,” Phys. Chem. Chem.Phys., vol. 2, no. 8, pp. 1685–1692, 2000.

[78] H. Sze, X. Li, and M. G. Palmgren, “Energization of plant cell mem-branes by H+-pumping ATPases. Regulation and biosynthesis,” PlantCell, vol. 11, no. 4, pp. 677–690, Apr. 1999.

[79] D. Foxx and E. E. Kalu, “Amperometric biosensor based on thermally ac-tivated polymer-stabilized metal nanoparticles,” Electrochem. Commun.,vol. 9, no. 4, pp. 584–590, Apr. 2007.

[80] C. M. Moore, S. Hackman, T. Brennan, and S. D. Minteer, “Effect ofmixture casting phosphonium salts with Nafion on the proton exchangecapacity and mass transport through the membranes,” J. Membr. Sci.,vol. 254, no. 1/2, pp. 63–70, Jun. 2005.

[81] J. C. Biffinger, R. Ray, B. Little, and B. R. Ringeisen, “Diversifyingbiological fuel cell designs by use of nanoporous filters,” Environ. Sci.Technol., vol. 41, no. 4, pp. 1444–1449, Feb. 2007.

[82] Y. Mohana, S. Manoj Muthu Kumara, and D. Das, “Electricity genera-tion using microbial fuel cells,” Int. J. Hydrogen Energy, vol. 33, no. 1,pp. 423–426, 2008.

[83] A. B. Lethninger, Principle of Biochemistry. New York: Worth, 1982,pp. 467–505.

[84] J. K. Dormer and R. Z. Gan, “Implantable electronic otologic devices:State of the art, otolaryngol,” Clin. North Amer., vol. 34, no. 2, pp. 289–297, 2001.

[85] Y. Xia and G. M. Whitesides, “Soft lithography,” Annu. Rev. Mater. Sci.,vol. 27, pp. 153–184, 1998.

[86] J. Larminie and A. Dicks, Fuel Cell Systems Explained, 2nd ed.Chichester, U.K.: Wiley, 2003, pp. 26–73.

[87] S. Wilkinson, “‘Gastrobots’—Benefits and challenges of microbial fuelcells in foodpowered robot applications,” J. Auton. Robots, vol. 9, no. 2,pp. 99–111, Sep. 2000.

[88] M. DuBois, K. A. Gilles, J. K. Hamilton, P. A. Rebers, and F. Smith,“Colorimetric method for determination of sugars and related sub-stances,” Anal. Chem., vol. 28, pp. 350–356, 1956.

[89] G. C. Johnston, C. W. Ehrhardt, A. Lorincz, and B. L. Carter, “Regu-lation of cell size in the yeast Saccharomyces cerevisiae,” J. Bacteriol.,vol. 137, no. 1, pp. 1–5, Jan. 1979.

[90] F. M. Harold, “To shape a cell: An inquiry into the causes of morphogen-esis of microorganisms,” Microbiol. Rev., vol. 54, no. 4, pp. 381–431,Dec. 1990.

[91] G. M. Walker, Yeast Physiology and Biotechnology. New York: Wiley,1998.

[92] U. Theobald, W. Mailinger, M. Baltes, M. Rizzi, and M. Reuss, “Invivo analysis of metabolic dynamics in Saccharomyces cerevisiae:I. Experimental observations,” Biotechnol. Bioeng., vol. 55, no. 2,pp. 305–316, Mar. 2000.

[93] C. Basheer, S. Swaminathan, H. K. Lee, and S. Valiyaveettil, “Develop-ment and application of a simple capillary-microreactor for oxidation ofglucose with a porous gold catalyst,” Chem. Commun., vol. 21, no. 3,pp. 409–410, Jan. 2005.

[94] D. R. Crow, Principles and Applications of Electrochemistry, 3rd ed.London, U.K.: Chapman & Hall, 1988, pp. 83–164.

[95] S. J. Lin, E. Ford, M. Haigis, G. Liszt, and L. Guarente, “Calorie restric-tion extends yeast life span by lowering the level of NADH,” Genes Dev.,vol. 18, no. 1, pp. 12–16, 2004.

[96] P. W. Atkins, Physical Chemistry, 6th ed. New York: Freeman, 1998,pp. 173–175.

[97] American Public Health Association, American Water Works Associ-ation, Water Pollution Control Federation, Water Environment Federa-tion, Standard Methods for the Examination of Water and Wastewater,12th ed. New York: APHA, 1965, pp. 408–410.

[98] A. Kumlanghan, J. Liu, P. Thavarungkul, P. Kanatharanab, andB. Mattiasson, “Microbial fuel cell-based biosensor for fast analysis ofbiodegradable organic matter,” Biosens. Bioelectron., vol. 22, no. 12,pp. 2939–2944, Jun. 2007.

[99] T. Catal, K. Li, H. Bermek, and H. Liu, “Electricity production fromtwelve monosaccharides using microbial fuel cells,” J. Power Sources,vol. 175, pp. 196–200, 2008.

[100] K. Kovárová-Kovar and T. Egli, “Growth kinetics of suspended micro-bial cells: From single-substrate-controlled growth to mixed-substratekinetics,” Microbiol. Mol. Biol. Rev., vol. 62, no. 3, pp. 646–666,1998.

[101] A. K. Manohar, O. Bretschger, H. Nealson, and F. Mansfeld, “The useof Electrochemical Impedance Spectroscopy (EIS) in the evaluation of



the performance of microbial fuel cells,” Bioelectrochemistry, vol. 72,pp. 149–154, 2008.

[102] R. Gifford, J. J. Kehoe, S. L. Barnes, B. A. Kornilayev, M. A. Alterman,and G. S. Wilson, “Protein interactions with subcutaneously implantedbiosensors,” Biomaterials, vol. 28, no. 12, pp. 2587–2598, Apr. 2006.

[103] N. Wisniewski and M. Reichert, “Methods for reducing biosensor mem-brane biofouling,” Colloids Surf. B, Biointerfaces, vol. 18, no. 3/4,pp. 197–219, Oct. 2000.

[104] D. P. Jones, J. L. Carlson, V. C. Mody, J. Cai, M. J. Lynn, andP. Sternberg, “Redox state of glutathione in human plasma,” Free Radic.Biol. Med., vol. 28, no. 4, pp. 625–635, Feb. 2000.

[105] D. P. Jones, V. C. Mody, J. L. Carlson, M. J. Lynn, and P. Sternberg, “Re-dox analysis of human plasma allows separation of pro-oxidant eventsof aging from decline in antioxidant defenses,” Free Radic. Biol. Med.,vol. 33, no. 9, pp. 1290–1300, Nov. 2002.

[106] J. Menicucci, H. Beyenal, E. Marsili, R. A. Veluchamy, G. Demir, andZ. Lewandowski, “Procedure for determining maximum sustainablepower generated by microbial fuel cells,” Environ. Sci. Technol., vol. 40,no. 3, pp. 1062–1068, Feb. 2006.

[107] J. Toffaletti, Special Topics in Diagnostic Testing. Washington, DC:AACC Press, 2001, pp. 53–70.

Chin-Pang-Billy Siu (S’06) received the B.S. de-gree in applied physics from the Hong Kong BaptistUniversity, Hong Kong, in 1996 and the B.Eng. andM.Phil. degrees in mechanical engineering from theHong Kong University of Science and Technology,Hong Kong, in 1999 and 2004, respectively. He iscurrently working toward the Ph.D. degree at the De-partment of Mechanical Engineering, The Universityof British Columbia, Vancouver, BC, Canada.

He was a Project Engineer with OEM householdelectronics development and a Consulting Engineer

who provided I-beam foundation and hoisting machinery designs and consult-ing services for the Hong Kong government and private companies. His currentresearch activities include MEMS microbial fuel cells and scanning confocalmicrolenses for Raman spectroscopy.

Mr. Siu was the recipient of the 2006 Example of IC’s manufactured CMP forthe “Magnetically actuated scanning microlens for NIR Raman spectroscopy,”and the 2008 Strategic Microelectronics Council of Information Technol-ogy Association of Canada Industrial Collaboration Awards for research in“A MetalMUMPs fabrication process for dual-axes scanning microlens.”

Mu Chiao(M’04)received the B.S. and M.S. degreesfrom National Taiwan University, Taipei, Taiwan,in 1996, and the Ph.D. degree in mechanical engi-neering from the University of California, Berkeley,in 2002.

From 2002 to 2003, he was with the Berkeley Sen-sor and Actuator Center, University of California,Berkeley, as a Postdoctoral Research Fellow. Since2003, he has been with the Department of Mechani-cal Engineering, The University of British Columbia,Vancouver, BC, Canada, where he is currently an

Associate Professor. His current research interests include design and fabri-cation of MEMS and nanodevices for biomedical applications.

Dr. Chiao is supported by the Canada Research Chairs, Tier 2 program.


Documents

A Micro Fabricated PDMS Microbial Fuel Cell 2008