Article pubs.acs.org/Langmuir
Microcontact Imprinting of Algae for Biofuel Systems: The Effects of the Polymer Concentration Mei-Hwa Lee,† James L. Thomas,‡ Ming-Yuan Lai,§ Ching-Ping Shih,§ and Hung-Yin Lin*,§ †
Department of Materials Science and Engineering, I-Shou University, Kaohsiung 84001, Taiwan Department of Physics and Astronomy, University of New Mexico, Albuquerque, New Mexico 87131, United States § Department of Chemical and Materials Engineering, National University of Kaohsiung, Kaohsiung 81148, Taiwan ‡
ABSTRACT: Microcontact imprinting of cells often involves the deposition of a polymer solution onto a monolayer cell stamp, followed by solvent evaporation. Thus, the concentration of the polymer may play an important role in the final morphology and efficacy of the imprinted film. In this work, various concentrations of poly(ethylene-co-vinyl alcohol) (EVAL) were dissolved in dimethyl sulfoxide (DMSO) for the microcontact imprinting of algae on an electrode. Scanning electron microscopy and fluorescence spectrometry were used to characterize the surface morphology and recognition capacity of algae to the algae-imprinted cavities. The readsorption of algae onto algae-imprinted EVAL thin films was quantified to obtain the EVAL concentration that maximized algal binding. Finally, the power and current density of an algal biofuel cell with the algae-imprinted EVAL-coated electrode were measured and found to be approximately double those of such a cell with a Pt/indium tin oxide (ITO)/poly(ethylene terephthalate) (PET) electrode.
■
INTRODUCTION There is growing interest in microimprinting of cells into various polymer matrices, because the cellular microenvironment has been shown to influence cell behavior and gene expression.1−6 Imprints of differentiated cells can cause stem cells to differentiate into the imprinted phenotype.2 Cellular imprinting has been used to identify the growth stages of different bacteria.7,8 Osteosarcoma cells that were cultured for longer (14 days versus 4 h) before imprinting gave, on average, rougher and deeper imprinted cavities;6 cells that grew on those imprinted surfaces exhibited higher cellular viability and activity and may have had altered gene expression as well. Note that, to prevent damage to cellular membranes during the imprinting process, special protocols must be followed (e.g., cells were fixed) before the polymerization of cell-imprinted polymers (CIPs).1,2,5,7 The three main methods for preparing cell-imprinted polymers are as follows: (1) surface imprinting with celladsorbed stamps, which are covered with monomer solutions, which polymerize to form, for example, polyurethane (PU),8−11 (2) culturing of cells on Petri dishes, followed by cell imprinting into a silicone [such as polydimethylsiloxane (PDMS)],1,2,5−7,12 and (3) cell imprinting into colloids, formed by a mixture of tetraethyl orthosilicate (TEOS) and a catalyst (such ammonia).4,13−15 Over the past decade, Dickert’s group has employed the surface-imprinting method (1) to prepare cell-imprinted receptors on a quartz crystal microbalance (QCM), sensing chips for the mass sensing of yeasts, viruses, and erythrocytes.8−11,15 A similar method was also recently © 2014 American Chemical Society
applied to the selective identification of macrophages and cancer cells.12 More recently, soft lithography (method 2) has been used to produce a negative replica of cells that were fixed and covered with pre-polymer solution (such as PDMS and a cross-linker).1,2,5−7,12 Zare’s group subsequently used the PDMS replica to separate and sort bacteria.1,5,7 Jeon and Kim6 used it to elucidate the relationship between surface patterns and cellular activities (of, for example, MG63 osteoblast-like cells), and Mahmoudi et al. used such imprints to direct the differentiation of stem cells.2 In this work, the effect of the concentration of the imprinting poly(ethylene-co-vinyl alcohol) (EVAL) solution [in dimethyl sulfoxide (DMSO)] was studied. A glass slide with a monolayer of adsorbed algae was employed as the cell-imprinting stamp; several EVAL concentrations in DMSO were used to form algae-imprinted EVAL thin films (AIPs) by solvent evaporation. The distribution of pore sizes, the film thickness, and the readsorption of the AIPs were determined. The power output of algal biofuel cells with an AIP-coated anode was then measured for the comparison of the imprinting polymer concentration effect.
■
EXPERIMENTAL SECTION
EVAL containing ethylene of 38 mol % was purchased from Scientific Polymer Products (Ontario, NY). Sodium dodecyl sulfate (SDS), Received: August 5, 2014 Revised: October 21, 2014 Published: October 30, 2014 14014
dx.doi.org/10.1021/la5031119 | Langmuir 2014, 30, 14014−14020
Langmuir
Article
Scheme 1. Preparation and EVAL Concentration Affect the Output of the Microcontact Algae-Imprinted Pt/ITO/PET Electrode for Biofuel Cells
DMSO, and potassium hydrogen phosphate were from J.T.Baker (ACS grade, Phillipsburg, NJ). Chlamydomonas reinhardtii was a generous gift by Professor Chung-Kuang Lu at the National Museum of Marine Biology and Aquarium (Pingtung, Taiwan). Sodium nitrate was from Wako Pure Chemical, Ltd. (Osaka, Japan). Calcium chloride dihydrate and potassium dihydrogen phosphate were from Riedel-de Haën Co. (Germany). Sodium chloride and magnesium sulfate heptahydrate were from Sigma-Aldrich Co. (St. Louis, MO). Proteose pertone was from Fluka Biochemika (Buchs, Switzerland). Nafion perfluorosulfonic acid (PFSA) membrane N117 was from DuPont Fuel Cells (Wilmington, DE). A 3.0 × 3.0 cm2 Nafion 117 film was used as the proton exchange membrane (PEM). The culture medium for C. reinhardtii contained 2.94 mM sodium nitrate, 0.17 mM calcium chloride dehydrate, 0.30 mM magnesium sulfate heptahydrate, 0.43 mM potassium hydrogen phosphate, 1.29 mM potassium dihydrogen phosphate, 0.43 mM sodium chloride, and 0.1% proteose pertone. All chemicals were used as received, unless otherwise mentioned. A indium tin oxide (ITO) coated with poly(ethylene terephthalate) (PET) thin film was cut into the size of 2.5 × 3.0 cm2, cleaned with detergent, and then dried in an oven at 60 °C. The ITO/PET thin film was sputtered with platinum at 10 mA for 300 s with an ion sputter coater (Hitachi E-1045). Materials used for electrodes have included carbon paper,16 carbon cloth,17,18 carbon mesh,19 graphite rods,20 stainless-steel plate,21 and platinum mesh.22 In this work, we selected a sputter-coated platinum electrode to enhance the conductivity; the use of ITO may have the same advantage. Moreover, the electrode area can be well-defined by sputtering on a planar plastic thin film for microcontact imprinting and compared to our previous work.3 As shown in Scheme 1, the preparation of an algae-imprinted EVAL thin film by microcontact imprinting23 included the following steps: adsorption of algae in a 2 mL (1 × 106, 5 × 106, 1 × 107, and 2 × 107) of cells/mL of algae solution for 30 min using glass slides in the size of 3.0 × 3.0 cm2 to cover the electrode area. The algae stamps were previously cleaned with isopropanol, deionized water, ethanol, and deionized water in 55 °C for 30 min under sonication. Moisture on the stamps could be dried under very gentle nitrogen blowing. Then, various concentrations of EVAL solutions from 1.0 to 10.0 wt % in DMSO were applied onto the algae stamps, covered with the Pt/ITO/ PET electrode, and dried in a vacuum oven at 60 °C for 3 h to remove solvents. Finally, the algae-imprinted EVAL Pt/ITO/PET electrode
was peeled off and washed with deionized water and phosphatebuffered saline (PBS) 3 times and 10 min each time. Algae- and non-imprinted polymers were freeze-dried before examination by a scanning electron microscope (Hitachi S4700, Hitachi High-Technologies Co., Tokyo, Japan). The image analysis used software ImageJ (http://imagej.nih.gov/ij/index.html) and setting the scale using the scanning electron microscopy (SEM) scale bar. “Freehand selection” was used to manually surround each cavity for measurement of its area and effective diameters. The pore diameter distributions and their average were calculated from the measured pore area, and the pore densities were estimated from the number of pores in the entire image, divided by the image area. The true cross-sections of samples were obtaining by placing into liquid nitrogen (caution: do not handle liquid nitrogen until you have read the cautionary notes) until it stops bubbling. The material was removed and then cracked using heavy-duty tweezers. The adsorption of algae to the algae- and non-imprinted polymer films was examined by immersion into 2 mL algae solution (107 cells/mL) for 60 min and then measuring the algae concentration in the solution with a fluorescence spectrophotometer (F-7000, Hitachi Co., Japan), with excitation and emission wavelengths of 485 and 685 nm, respectively. The algae cell numbers can be calibrated with fluorescence intensity. All parts of the fuel cells were sterilized in an autoclave and irradiated under ultraviolet (UV, 15 W, G15T8, Philips, Amsterdam, Netherlands; it has an UV output of 4.9 W at a wavelength of 254 nm and is used in sterilizing applications) in a laminar flow hood overnight before assembling. After adding 250 mL of PBS to the cathode cell, the algae culture medium (without the addition of magnesium sulfate heptahydrate) was added to the anode cell. The anode cell was then purged with nitrogen gas to enhance hydrogen production. Finally, a platinum wire 5 cm long and the algae-imprinted EVAL-coated Pt/ ITO/PET electrode were used as the cathode and anode, respectively. A potentiostat (model 608-1A, CH Instruments, Inc., Austin, TX) was employed to measure the current output by an amperometric I−t curve. For illumination during the power measurements, three white light fluorescent lamps (14 W, FH14D-EX/T, China Electric Manufacturing Corporation, Taiwan) at a distance of 30 cm were used. The illumination causes the production of hydrogen; H2producing illuminated C. reinhardtii colonies were observed by Ghirardi et al.24 The initial voltages were decreased from the maximum voltage [i.e., open circuit voltage (OCV)] output by 14015
dx.doi.org/10.1021/la5031119 | Langmuir 2014, 30, 14014−14020
Langmuir
Article
Figure 1. (a) Preparation of the algae stamp for the microcontact-imprinted polymers. (b−e) Optical images of the algae adsorption under 1.0 × 106, 5.0 × 106, 1.0 × 107, and 2.0 × 107 cells/mL of algae solution for 60 min. −0.05 V/step. Although the best technique for fuel cell measurements is to force a cell current (galvanostatic control) and measure the resulting voltage, in electrochemistry studies, it is common practice to fix the potential of a cell (potentiostatic control) and measure the resulting cell current.25 Power density (P = VI/A) was calculated from the measured voltage (V), current (I = V/R), and surface area of the anode electrode (A).26 The polarization and power curves were the plots of current versus voltage and power output, respectively.
(Figure 1). In adhesion from 106 cells/mL (Figure 1b), algae were present as isolated cells or in groups of only a few algal cells. At higher algae concentrations, the algae aggregate sizes increased, forming an infinite network at 107 cells/mL (Figure 1d) and a confluent monolayer at 2 × 107 cells/mL (Figure 1e). For further work here, we used the highest concentration algal suspension; as will be seen, this confluent monolayer stamp gave better performance (e.g., 5-fold algae adsorption) than we obtained previously.3 Figure 2 presents SEM images of stamps that were made by microcontact-imprinting 1.0−10.0 wt % EVAL (containing 38 mol % ethylene) using confluent algal cells as a stamp (2 × 107 cell/mL). EVAL concentrations of 1.0, 3.0, 5.0, 7.0, and 10.0 wt % were used. The right-hand column of Figure 2 shows histograms of the sizes of the imprinted cavities. At the lowest imprinting EVAL concentrations (Figure 2a), 1.0 and 3.0 wt %, many imprinted cavities were incomplete. The shape of the imprinted algae is clearer at EVAL concentrations of 5.0 and 7.0 wt %. Finally, the polymer may completely fill the spaces between the algae at an EVAL concentration of 10.0 wt %
■
RESULTS AND DISCUSSION The density of templates (particularly template molecules) may play an important role in the recognition of microcontactimprinted polymers. In particular, when preparing molecularly imprinted polymers, monolayer or sub-monolayer template densities are typically used, because multi-layer adsorption of template molecules on the stamp may reduce the ability of the imprinted cavities to recognize a target molecule. In an earlier study, we employed an algal stamp formed by allowing algae from a suspension of 107 cells/mL to adhere to a glass slide for 30 min.3 In this study, we observed algal adhesion using suspension concentrations from 1 × 106 to 2 × 107 cells/mL 14016
dx.doi.org/10.1021/la5031119 | Langmuir 2014, 30, 14014−14020
Langmuir
Article
Figure 2. (a and b) SEM images and (c) imprinted cavity distributions of the algae-imprinted EVAL (containing 38 mol % ethylene) thin films with the EVAL concentrations from 1.0 to 10.0 wt %.
Table 1. Average Imprinted Area and Density of Cavities on AIPs Using Different Concentrations of EVALs Calculated from SEM Images Shown in Figure 2 EVAL concentration (wt %) imprinted cavities 2
average area (μm ) average density (×103 cells/mm2)
1.0
3.0
5.0
7.0
10.0
59.9 ± 19.2 4.2 ± 0.3
79.6 ± 17.1 4.8 ± 1.6
87.1 ± 19.8 6.0 ± 0.5
90.6 ± 17.2 6.1 ± 1.3
78.4 ± 13.4 6.1 ± 0.6
concentration increased from 1.0 to 7.0 wt % in Figure 2c. The average pore diameters were 8.61 ± 1.51, 10.01 ± 1.07, 10.46 ±
EVAL. The peaks of the alga-imprinted pore size distributions increased from approximately 8 to 10−12 μm as the EVAL 14017
dx.doi.org/10.1021/la5031119 | Langmuir 2014, 30, 14014−14020
Langmuir
Article
1.20, 10.69 ± 1.07, and 9.96 ± 0.86 μm, respectively. Table 1 presents the mean area and density of the imprinted cavities. The average densities of those cavities increased from 4.2 ± 0.3 to 6.1 ± 0.60 × 103 cells/mm2 when the imprinting EVAL concentration increased from 1.0 to 7.0 wt %. Figure 3 shows the thicknesses of the algae-imprinted EVAL thin films, which increased linearly with the EVAL concen-
Figure 4. Saturated adsorption with different algae solution concentrations on 38 mol % ethylene imprinted and non-imprinted EVAL films. The EVAL concentrations applied were 5.0 or 7.0 wt %.
(10%), although it gave the thickest films, did not give the highest saturated absorption of algae. The saturating adsorption on 10 wt % EVAL was comparable to that obtained with 5 wt %. This result suggests that the recognition of algae is mainly via surface proteins or complexes, which were imprinted on the polymer surface; if recognition were via the whole cell topography, one would have expected that a thicker film (i.e., comparable to the cell diameter) would have worked better. Algal biofuel cells with an algae concentration of 1.0 × 107 cells/mL were assembled to evaluate the power output of algaebinding anodes imprinted using different concentrations of EVAL. As presented in Figure 5a, the output voltage when algae-imprinted EVAL and a bare Pt/ITO/PET electrode were used reveals the existence of a latent period before the initial output. After the latent period, the algal fuel cells were able to operate continuously for at least 100 h (unlike microbial fuel cells). Figure 5b plots the OCVs of algal biofuel cells using AIPcoated, NIP-coated, and bare electrodes. The OCV is around 0.63 ± 0.01 and 0.53 ± 0.02 V for AIPs and NIPs, respectively, indicating that the electron-transfer efficiency from algae exceeds that of a bare or NIP-coated electrode. Biological redox potentials of electron donors and acceptors28 at pH 7 for H+/H2 and O2/H2O are −0.41 and +0.82 V versus a standard hydrogen electrode (SHE), respectively. Figure 5c plots the measured polarization behavior of the algae fuel cells. The output voltages decreased as the loading current density increased. The use of ITO-coated PET greatly increased the current over that in our earlier work.3 When the algaeimprinted 7 wt % EVAL-coated Pt/ITO/PET electrode was used, the output power density was approximately 1.65 ± 0.16 mW/m2 at a current density of 9.10 mA/m2, which was about double and triple the power obtained using bare and NIPcoated Pt/ITO/PET electrodes, as shown in Figure 5d. To make an estimate of the variability, at least two 7 wt % electrodes were made and the standard deviation in performance was measured. We expect that the deviations for other compositions are similar.
Figure 3. (a−c) SEM cross-section images of the algae-imprinted EVAL (containing 38 mol % ethylene) thin films with the EVAL concentrations of (a) 1.0, (b) 5.0, and (c) 10.0 wt %. (d) Thickness of the algae-imprinted EVAL thin film increased with the EVAL concentration.
trations. Interestingly, at the highest EVAL concentration, the film thickness was 5.4 ± 0.2 μm or about half the diameter of an algal cell (ca. 10 μm from optical microscopy images). The adsorption of algae on imprinted or non-imprinted EVAL (38 mol % ethylene) thin films saturated at algae concentrations of about 2.0 × 107 cells/mL, as displayed in Figure 4. The readsorption results were fit by the Hill equation, giving the maximum adsorptions for 5.0 and 7.0 wt % of 4 × 106 and 5.0 × 106 cells/cm2, respectively. The standard deviation in those adsorption measurements is approximately ±5%. A higher EVAL concentration (10.0 wt %) did not provide a higher recognition capacity (saturating at approximately 4.4 × 106 cells/cm2). For non-imprinted polymers (NIPs), the average maximum adsorption was 2.4 × 106 cells/cm2. In all cases (including NIPs), the Hill coefficient, which quantifies cooperativity in binding, was 2−3. The Hill equation is equivalent to the Langmuir equation when the Hill coefficient = 1, which would have indicated non-cooperative binding. It seems quite unlikely that conformational changes in the film lead to cooperativity in binding (especially with non-imprinted films); rather, the Hill coefficient likely reflects increasing algal aggregation in solution as the concentration is increased, leading to binding of increasing algal dimers, trimers, and larger aggregates.27 Interestingly, the highest EVAL concentration 14018
dx.doi.org/10.1021/la5031119 | Langmuir 2014, 30, 14014−14020
Langmuir
Article
Figure 5. (a) Time course measurements of the OCV of algae fuel cells using algae-imprinted EVAL-coated and bare Pt/ITO/PET electrodes as the anode, (b) OCV changes with different EVAL concentrations applied for AIP- and NIP-coated anodes in biofuel cells, (c) polarization behavior of the algal fuel cells, with voltage (open symbols) and power (filled symbols) changing with the loading of different resistances using AIP electrodes made with various polymer concentrations, and (d) maximum power related to EVAL concentrations used for microcontact imprinting.
■
The 7 wt % EVAL films gave the better adsorption than 5 wt % (Figure 4) and gave the best electrochemical performance, even though both films appeared to have nearly the same density of cavities. We hypothesize that the thinner film is unable to form cavities of the same quality or affinity, while thicker films offer no further improvement in binding and actually hinder H2 transport.
■
AUTHOR INFORMATION
Corresponding Author
*Telephone: +886-7-591-9455 and/or +886-912-178-751. Email: [email protected] and/or [email protected]. Notes
The authors declare no competing financial interest.
■
ACKNOWLEDGMENTS The authors acknowledge the National Science Council of the Republic of China (ROC), Taiwan, for financially supporting this research under Contracts NSC 102-2220-E-390-001 and NSC 102-2220-E-006-004 and the Ministry of Science and Technology of the ROC under Contracts MOST 103-2220-E390-001 and MOST 103-2220-E-006-007.
CONCLUSION
The present work reveals that algae-imprinted polymers made by 7.0 wt % EVAL that contains 38 mol % ethylene were better able to recognize algae than other polymer concentrations. In this specific polymer concentration, the average pore diameter of the algae-imprinted polymer was 10.69 ± 1.07 μm, to provide the highest saturated absorption of algae. An electrochemical performance comparison between the algaeimprinted EVAL-coated and bare Pt/ITO/PET electrodes indicated that the former achieved a power output of 1.65± 0.16 mW/m2 at a current density of 9.10 mA/m2; this power is substantially higher than that obtained with bare or NIP-coated algal electrodes. These results suggest that the polymer microenvironment may increase the metabolic activity of algae.
■
REFERENCES
(1) Ren, K.; Banaei, N.; Zare, R. N. Sorting inactivated cells using cell-imprinted polymer thin films. ACS Nano 2013, 7, 6031−6036. (2) Mahmoudi, M.; Bonakdar, S.; Shokrgozar, M. A.; Aghaverdi, H.; Hartmann, R.; Pick, A.; Witte, G.; Parak, W. J. Cell-imprinted substrates direct the fate of stem cells. ACS Nano 2013, 7, 8379−8384. (3) Chen, W.-J.; Lee, M.-H.; Thomas, J. L.; Lu, P.-H.; Li, M.-H.; Lin, H.-Y. Microcontact imprinting of algae on poly(ethylene-co-vinyl
14019
dx.doi.org/10.1021/la5031119 | Langmuir 2014, 30, 14014−14020
Langmuir
Article
alcohol) for biofuel cells. ACS Appl. Mater. Interfaces 2013, 5, 11123− 11128. (4) Borovicka, J.; Stoyanov, S. D.; Paunov, V. N. Shape recognition of microbial cells by colloidal cell imprints. Nanoscale 2013, 5, 8560− 8568. (5) Ren, K.; Zare, R. N. Chemical recognition in cell-imprinted polymers. ACS Nano 2012, 6, 4314−4318. (6) Jeon, H.; Kim, G. Effects of a cell-imprinted poly(dimethylsiloxane) surface on the cellular activities of MG63 osteoblast-like cells: Preparation of a patterned surface, surface characterization, and bone mineralization. Langmuir 2012, 28, 13423−13430. (7) Schirhagl, R.; Hall, E. W.; Fuereder, I.; Zare, R. N. Separation of bacteria with imprinted polymeric films. Analyst 2012, 137, 1495− 1499. (8) Dickert, F. L.; Hayden, O.; Halikias, K. P. Synthetic receptors as sensor coatings for molecules and living cells. Analyst 2001, 126, 766− 771. (9) Hayden, O.; Bindeus, R.; Haderspöck, C.; Mann, K.-J.; Wirl, B.; Dickert, F. L. Mass-sensitive detection of cells, viruses and enzymes with artificial receptors. Sens. Actuators, B 2003, 91, 316−319. (10) Hayden, O.; Lieberzeit, P. A.; Blaas, D.; Dickert, F. L. Artificial antibodies for bioanalyte detectionSensing viruses and proteins. Adv. Funct. Mater. 2006, 16, 1269−1278. (11) Jenik, M.; Schirhagl, R.; Schirk, C.; Hayden, O.; Lieberzeit, P.; Blaas, D.; Paul, G.; Dickert, F. L. Sensing picornaviruses using molecular imprinting techniques on a quartz crystal microbalance. Anal. Chem. 2009, 81, 5320−5326. (12) Eersels, K.; van Grinsven, B.; Ethirajan, A.; Timmermans, S.; Jiménez Monroy, K. L.; Bogie, J. F. J.; Punniyakoti, S.; Vandenryt, T.; Hendriks, J. J. A.; Cleij, T. J.; Daemen, M. J. A. P.; Somers, V.; De Ceuninck, W.; Wagner, P. Selective identification of macrophages and cancer cells based on thermal transport through surface-imprinted polymer layers. ACS Appl. Mater. Interfaces 2013, 5, 7258−7267. (13) Borovička, J.; Metheringham, W. J.; Madden, L. A.; Walton, C. D.; Stoyanov, S. D.; Paunov, V. N. Photothermal colloid antibodies for shape-selective recognition and killing of microorganisms. J. Am. Chem. Soc. 2013, 135, 5282−5285. (14) Cohen, T.; Starosvetsky, J.; Cheruti, U.; Armon, R. Whole cell imprinting in sol−gel thin films for bacterial recognition in liquids: Macromolecular fingerprinting. Int. J. Mol. Sci. 2010, 11, 1236−1252. (15) Dickert, F. L.; Hayden, O. Bioimprinting of polymers and sol− gel phases. Selective detection of yeasts with imprinted polymers. Anal. Chem. 2002, 74, 1302−1306. (16) Luo, Y.; Liu, G.; Zhang, R.; Zhang, C. Power generation from furfural using the microbial fuel cell. J. Power Sources 2010, 195, 190− 194. (17) Catal, T.; Li, K.; Bermek, H.; Liu, H. Electricity production from twelve monosaccharides using microbial fuel cells. J. Power Sources 2008, 175, 196−200. (18) Rezaei, F.; Xing, D.; Wagner, R.; Regan, J. M.; Richard, T. L.; Logan, B. E. Simultaneous cellulose degradation and electricity production by Enterobacter cloacae in a microbial fuel cell. Appl. Environ. Microbiol. 2009, 75, 3673−3678. (19) Wang, X.; Cheng, S.; Feng, Y.; Merrill, M. D.; Saito, T.; Logan, B. E. Use of carbon mesh anodes and the effect of different pretreatment methods on power production in microbial fuel cells. Environ. Sci. Technol. 2009, 43, 6870−6874. (20) Patil, S. A.; Surakasi, V. P.; Koul, S.; Ijmulwar, S.; Vivek, A.; Shouche, Y. S.; Kapadnis, B. P. Electricity generation using chocolate industry wastewater and its treatment in activated sludge based microbial fuel cell and analysis of developed microbial community in the anode chamber. Bioresour. Technol. 2009, 100, 5132−5139. (21) Dumas, C.; Mollica, A.; Féron, D.; Basséguy, R.; Etcheverry, L.; Bergel, A. Marine microbial fuel cell: Use of stainless steel electrodes as anode and cathode materials. Electrochim. Acta 2007, 53, 468−473. (22) Rosenbaum, M.; Schröder, U.; Scholz, F. Utilizing the green alga Chlamydomonas reinhardtii for microbial electricity generation: A living solar cell. Appl. Microbiol. Biotechnol. 2005, 68, 753−756.
(23) Lin, H.-Y.; Hsu, C.-Y.; Thomas, J. L.; Wang, S.-E.; Chen, H.-C.; Chou, T.-C. The microcontact imprinting of proteins: The effect of cross-linking monomers for lysozyme, ribonuclease A and myoglobin. Biosens. Bioelectron. 2006, 22, 534−543. (24) Ghirardi, M. L.; Zhang, L.; Lee, J. W.; Flynn, T.; Seibert, M.; Greenbaum, E.; Melis, A. Microalgae: A green source of renewable H2. Trends Biotechnol. 2000, 18, 506−511. (25) Niemann, J. Unraveling fuel cell electrical measurements. Fuel Cell Mag. 2005, April/May, 26−31. (26) Min, B.; Logan, B. E. Continuous electricity generation from domestic wastewater and organic substrates in a flat plate microbial fuel cell. Environ. Sci. Technol. 2004, 38, 5809−5814. (27) Ackleh, A. S.; Fitzpatrick, B. G. Modeling aggregation and growth processes in an algal population model: Analysis and computations. J. Math. Biol. 1997, 35, 480−502. (28) Thauer, R. K.; Jungermann, K.; Decker, K. Energy conservation in chemotrophic anaerobic bacteria. Bacteriol. Rev. 1977, 41, 100−180.
14020
dx.doi.org/10.1021/la5031119 | Langmuir 2014, 30, 14014−14020
Microcontact Imprinting of Algae for Biofuel Systems: The Effects of the Polymer Concentration Mei-Hwa Lee,† James L. Thomas,‡ Ming-Yuan Lai,§ Ching-Ping Shih,§ and Hung-Yin Lin*,§ †
Department of Materials Science and Engineering, I-Shou University, Kaohsiung 84001, Taiwan Department of Physics and Astronomy, University of New Mexico, Albuquerque, New Mexico 87131, United States § Department of Chemical and Materials Engineering, National University of Kaohsiung, Kaohsiung 81148, Taiwan ‡
ABSTRACT: Microcontact imprinting of cells often involves the deposition of a polymer solution onto a monolayer cell stamp, followed by solvent evaporation. Thus, the concentration of the polymer may play an important role in the final morphology and efficacy of the imprinted film. In this work, various concentrations of poly(ethylene-co-vinyl alcohol) (EVAL) were dissolved in dimethyl sulfoxide (DMSO) for the microcontact imprinting of algae on an electrode. Scanning electron microscopy and fluorescence spectrometry were used to characterize the surface morphology and recognition capacity of algae to the algae-imprinted cavities. The readsorption of algae onto algae-imprinted EVAL thin films was quantified to obtain the EVAL concentration that maximized algal binding. Finally, the power and current density of an algal biofuel cell with the algae-imprinted EVAL-coated electrode were measured and found to be approximately double those of such a cell with a Pt/indium tin oxide (ITO)/poly(ethylene terephthalate) (PET) electrode.
■
INTRODUCTION There is growing interest in microimprinting of cells into various polymer matrices, because the cellular microenvironment has been shown to influence cell behavior and gene expression.1−6 Imprints of differentiated cells can cause stem cells to differentiate into the imprinted phenotype.2 Cellular imprinting has been used to identify the growth stages of different bacteria.7,8 Osteosarcoma cells that were cultured for longer (14 days versus 4 h) before imprinting gave, on average, rougher and deeper imprinted cavities;6 cells that grew on those imprinted surfaces exhibited higher cellular viability and activity and may have had altered gene expression as well. Note that, to prevent damage to cellular membranes during the imprinting process, special protocols must be followed (e.g., cells were fixed) before the polymerization of cell-imprinted polymers (CIPs).1,2,5,7 The three main methods for preparing cell-imprinted polymers are as follows: (1) surface imprinting with celladsorbed stamps, which are covered with monomer solutions, which polymerize to form, for example, polyurethane (PU),8−11 (2) culturing of cells on Petri dishes, followed by cell imprinting into a silicone [such as polydimethylsiloxane (PDMS)],1,2,5−7,12 and (3) cell imprinting into colloids, formed by a mixture of tetraethyl orthosilicate (TEOS) and a catalyst (such ammonia).4,13−15 Over the past decade, Dickert’s group has employed the surface-imprinting method (1) to prepare cell-imprinted receptors on a quartz crystal microbalance (QCM), sensing chips for the mass sensing of yeasts, viruses, and erythrocytes.8−11,15 A similar method was also recently © 2014 American Chemical Society
applied to the selective identification of macrophages and cancer cells.12 More recently, soft lithography (method 2) has been used to produce a negative replica of cells that were fixed and covered with pre-polymer solution (such as PDMS and a cross-linker).1,2,5−7,12 Zare’s group subsequently used the PDMS replica to separate and sort bacteria.1,5,7 Jeon and Kim6 used it to elucidate the relationship between surface patterns and cellular activities (of, for example, MG63 osteoblast-like cells), and Mahmoudi et al. used such imprints to direct the differentiation of stem cells.2 In this work, the effect of the concentration of the imprinting poly(ethylene-co-vinyl alcohol) (EVAL) solution [in dimethyl sulfoxide (DMSO)] was studied. A glass slide with a monolayer of adsorbed algae was employed as the cell-imprinting stamp; several EVAL concentrations in DMSO were used to form algae-imprinted EVAL thin films (AIPs) by solvent evaporation. The distribution of pore sizes, the film thickness, and the readsorption of the AIPs were determined. The power output of algal biofuel cells with an AIP-coated anode was then measured for the comparison of the imprinting polymer concentration effect.
■
EXPERIMENTAL SECTION
EVAL containing ethylene of 38 mol % was purchased from Scientific Polymer Products (Ontario, NY). Sodium dodecyl sulfate (SDS), Received: August 5, 2014 Revised: October 21, 2014 Published: October 30, 2014 14014
dx.doi.org/10.1021/la5031119 | Langmuir 2014, 30, 14014−14020
Langmuir
Article
Scheme 1. Preparation and EVAL Concentration Affect the Output of the Microcontact Algae-Imprinted Pt/ITO/PET Electrode for Biofuel Cells
DMSO, and potassium hydrogen phosphate were from J.T.Baker (ACS grade, Phillipsburg, NJ). Chlamydomonas reinhardtii was a generous gift by Professor Chung-Kuang Lu at the National Museum of Marine Biology and Aquarium (Pingtung, Taiwan). Sodium nitrate was from Wako Pure Chemical, Ltd. (Osaka, Japan). Calcium chloride dihydrate and potassium dihydrogen phosphate were from Riedel-de Haën Co. (Germany). Sodium chloride and magnesium sulfate heptahydrate were from Sigma-Aldrich Co. (St. Louis, MO). Proteose pertone was from Fluka Biochemika (Buchs, Switzerland). Nafion perfluorosulfonic acid (PFSA) membrane N117 was from DuPont Fuel Cells (Wilmington, DE). A 3.0 × 3.0 cm2 Nafion 117 film was used as the proton exchange membrane (PEM). The culture medium for C. reinhardtii contained 2.94 mM sodium nitrate, 0.17 mM calcium chloride dehydrate, 0.30 mM magnesium sulfate heptahydrate, 0.43 mM potassium hydrogen phosphate, 1.29 mM potassium dihydrogen phosphate, 0.43 mM sodium chloride, and 0.1% proteose pertone. All chemicals were used as received, unless otherwise mentioned. A indium tin oxide (ITO) coated with poly(ethylene terephthalate) (PET) thin film was cut into the size of 2.5 × 3.0 cm2, cleaned with detergent, and then dried in an oven at 60 °C. The ITO/PET thin film was sputtered with platinum at 10 mA for 300 s with an ion sputter coater (Hitachi E-1045). Materials used for electrodes have included carbon paper,16 carbon cloth,17,18 carbon mesh,19 graphite rods,20 stainless-steel plate,21 and platinum mesh.22 In this work, we selected a sputter-coated platinum electrode to enhance the conductivity; the use of ITO may have the same advantage. Moreover, the electrode area can be well-defined by sputtering on a planar plastic thin film for microcontact imprinting and compared to our previous work.3 As shown in Scheme 1, the preparation of an algae-imprinted EVAL thin film by microcontact imprinting23 included the following steps: adsorption of algae in a 2 mL (1 × 106, 5 × 106, 1 × 107, and 2 × 107) of cells/mL of algae solution for 30 min using glass slides in the size of 3.0 × 3.0 cm2 to cover the electrode area. The algae stamps were previously cleaned with isopropanol, deionized water, ethanol, and deionized water in 55 °C for 30 min under sonication. Moisture on the stamps could be dried under very gentle nitrogen blowing. Then, various concentrations of EVAL solutions from 1.0 to 10.0 wt % in DMSO were applied onto the algae stamps, covered with the Pt/ITO/ PET electrode, and dried in a vacuum oven at 60 °C for 3 h to remove solvents. Finally, the algae-imprinted EVAL Pt/ITO/PET electrode
was peeled off and washed with deionized water and phosphatebuffered saline (PBS) 3 times and 10 min each time. Algae- and non-imprinted polymers were freeze-dried before examination by a scanning electron microscope (Hitachi S4700, Hitachi High-Technologies Co., Tokyo, Japan). The image analysis used software ImageJ (http://imagej.nih.gov/ij/index.html) and setting the scale using the scanning electron microscopy (SEM) scale bar. “Freehand selection” was used to manually surround each cavity for measurement of its area and effective diameters. The pore diameter distributions and their average were calculated from the measured pore area, and the pore densities were estimated from the number of pores in the entire image, divided by the image area. The true cross-sections of samples were obtaining by placing into liquid nitrogen (caution: do not handle liquid nitrogen until you have read the cautionary notes) until it stops bubbling. The material was removed and then cracked using heavy-duty tweezers. The adsorption of algae to the algae- and non-imprinted polymer films was examined by immersion into 2 mL algae solution (107 cells/mL) for 60 min and then measuring the algae concentration in the solution with a fluorescence spectrophotometer (F-7000, Hitachi Co., Japan), with excitation and emission wavelengths of 485 and 685 nm, respectively. The algae cell numbers can be calibrated with fluorescence intensity. All parts of the fuel cells were sterilized in an autoclave and irradiated under ultraviolet (UV, 15 W, G15T8, Philips, Amsterdam, Netherlands; it has an UV output of 4.9 W at a wavelength of 254 nm and is used in sterilizing applications) in a laminar flow hood overnight before assembling. After adding 250 mL of PBS to the cathode cell, the algae culture medium (without the addition of magnesium sulfate heptahydrate) was added to the anode cell. The anode cell was then purged with nitrogen gas to enhance hydrogen production. Finally, a platinum wire 5 cm long and the algae-imprinted EVAL-coated Pt/ ITO/PET electrode were used as the cathode and anode, respectively. A potentiostat (model 608-1A, CH Instruments, Inc., Austin, TX) was employed to measure the current output by an amperometric I−t curve. For illumination during the power measurements, three white light fluorescent lamps (14 W, FH14D-EX/T, China Electric Manufacturing Corporation, Taiwan) at a distance of 30 cm were used. The illumination causes the production of hydrogen; H2producing illuminated C. reinhardtii colonies were observed by Ghirardi et al.24 The initial voltages were decreased from the maximum voltage [i.e., open circuit voltage (OCV)] output by 14015
dx.doi.org/10.1021/la5031119 | Langmuir 2014, 30, 14014−14020
Langmuir
Article
Figure 1. (a) Preparation of the algae stamp for the microcontact-imprinted polymers. (b−e) Optical images of the algae adsorption under 1.0 × 106, 5.0 × 106, 1.0 × 107, and 2.0 × 107 cells/mL of algae solution for 60 min. −0.05 V/step. Although the best technique for fuel cell measurements is to force a cell current (galvanostatic control) and measure the resulting voltage, in electrochemistry studies, it is common practice to fix the potential of a cell (potentiostatic control) and measure the resulting cell current.25 Power density (P = VI/A) was calculated from the measured voltage (V), current (I = V/R), and surface area of the anode electrode (A).26 The polarization and power curves were the plots of current versus voltage and power output, respectively.
(Figure 1). In adhesion from 106 cells/mL (Figure 1b), algae were present as isolated cells or in groups of only a few algal cells. At higher algae concentrations, the algae aggregate sizes increased, forming an infinite network at 107 cells/mL (Figure 1d) and a confluent monolayer at 2 × 107 cells/mL (Figure 1e). For further work here, we used the highest concentration algal suspension; as will be seen, this confluent monolayer stamp gave better performance (e.g., 5-fold algae adsorption) than we obtained previously.3 Figure 2 presents SEM images of stamps that were made by microcontact-imprinting 1.0−10.0 wt % EVAL (containing 38 mol % ethylene) using confluent algal cells as a stamp (2 × 107 cell/mL). EVAL concentrations of 1.0, 3.0, 5.0, 7.0, and 10.0 wt % were used. The right-hand column of Figure 2 shows histograms of the sizes of the imprinted cavities. At the lowest imprinting EVAL concentrations (Figure 2a), 1.0 and 3.0 wt %, many imprinted cavities were incomplete. The shape of the imprinted algae is clearer at EVAL concentrations of 5.0 and 7.0 wt %. Finally, the polymer may completely fill the spaces between the algae at an EVAL concentration of 10.0 wt %
■
RESULTS AND DISCUSSION The density of templates (particularly template molecules) may play an important role in the recognition of microcontactimprinted polymers. In particular, when preparing molecularly imprinted polymers, monolayer or sub-monolayer template densities are typically used, because multi-layer adsorption of template molecules on the stamp may reduce the ability of the imprinted cavities to recognize a target molecule. In an earlier study, we employed an algal stamp formed by allowing algae from a suspension of 107 cells/mL to adhere to a glass slide for 30 min.3 In this study, we observed algal adhesion using suspension concentrations from 1 × 106 to 2 × 107 cells/mL 14016
dx.doi.org/10.1021/la5031119 | Langmuir 2014, 30, 14014−14020
Langmuir
Article
Figure 2. (a and b) SEM images and (c) imprinted cavity distributions of the algae-imprinted EVAL (containing 38 mol % ethylene) thin films with the EVAL concentrations from 1.0 to 10.0 wt %.
Table 1. Average Imprinted Area and Density of Cavities on AIPs Using Different Concentrations of EVALs Calculated from SEM Images Shown in Figure 2 EVAL concentration (wt %) imprinted cavities 2
average area (μm ) average density (×103 cells/mm2)
1.0
3.0
5.0
7.0
10.0
59.9 ± 19.2 4.2 ± 0.3
79.6 ± 17.1 4.8 ± 1.6
87.1 ± 19.8 6.0 ± 0.5
90.6 ± 17.2 6.1 ± 1.3
78.4 ± 13.4 6.1 ± 0.6
concentration increased from 1.0 to 7.0 wt % in Figure 2c. The average pore diameters were 8.61 ± 1.51, 10.01 ± 1.07, 10.46 ±
EVAL. The peaks of the alga-imprinted pore size distributions increased from approximately 8 to 10−12 μm as the EVAL 14017
dx.doi.org/10.1021/la5031119 | Langmuir 2014, 30, 14014−14020
Langmuir
Article
1.20, 10.69 ± 1.07, and 9.96 ± 0.86 μm, respectively. Table 1 presents the mean area and density of the imprinted cavities. The average densities of those cavities increased from 4.2 ± 0.3 to 6.1 ± 0.60 × 103 cells/mm2 when the imprinting EVAL concentration increased from 1.0 to 7.0 wt %. Figure 3 shows the thicknesses of the algae-imprinted EVAL thin films, which increased linearly with the EVAL concen-
Figure 4. Saturated adsorption with different algae solution concentrations on 38 mol % ethylene imprinted and non-imprinted EVAL films. The EVAL concentrations applied were 5.0 or 7.0 wt %.
(10%), although it gave the thickest films, did not give the highest saturated absorption of algae. The saturating adsorption on 10 wt % EVAL was comparable to that obtained with 5 wt %. This result suggests that the recognition of algae is mainly via surface proteins or complexes, which were imprinted on the polymer surface; if recognition were via the whole cell topography, one would have expected that a thicker film (i.e., comparable to the cell diameter) would have worked better. Algal biofuel cells with an algae concentration of 1.0 × 107 cells/mL were assembled to evaluate the power output of algaebinding anodes imprinted using different concentrations of EVAL. As presented in Figure 5a, the output voltage when algae-imprinted EVAL and a bare Pt/ITO/PET electrode were used reveals the existence of a latent period before the initial output. After the latent period, the algal fuel cells were able to operate continuously for at least 100 h (unlike microbial fuel cells). Figure 5b plots the OCVs of algal biofuel cells using AIPcoated, NIP-coated, and bare electrodes. The OCV is around 0.63 ± 0.01 and 0.53 ± 0.02 V for AIPs and NIPs, respectively, indicating that the electron-transfer efficiency from algae exceeds that of a bare or NIP-coated electrode. Biological redox potentials of electron donors and acceptors28 at pH 7 for H+/H2 and O2/H2O are −0.41 and +0.82 V versus a standard hydrogen electrode (SHE), respectively. Figure 5c plots the measured polarization behavior of the algae fuel cells. The output voltages decreased as the loading current density increased. The use of ITO-coated PET greatly increased the current over that in our earlier work.3 When the algaeimprinted 7 wt % EVAL-coated Pt/ITO/PET electrode was used, the output power density was approximately 1.65 ± 0.16 mW/m2 at a current density of 9.10 mA/m2, which was about double and triple the power obtained using bare and NIPcoated Pt/ITO/PET electrodes, as shown in Figure 5d. To make an estimate of the variability, at least two 7 wt % electrodes were made and the standard deviation in performance was measured. We expect that the deviations for other compositions are similar.
Figure 3. (a−c) SEM cross-section images of the algae-imprinted EVAL (containing 38 mol % ethylene) thin films with the EVAL concentrations of (a) 1.0, (b) 5.0, and (c) 10.0 wt %. (d) Thickness of the algae-imprinted EVAL thin film increased with the EVAL concentration.
trations. Interestingly, at the highest EVAL concentration, the film thickness was 5.4 ± 0.2 μm or about half the diameter of an algal cell (ca. 10 μm from optical microscopy images). The adsorption of algae on imprinted or non-imprinted EVAL (38 mol % ethylene) thin films saturated at algae concentrations of about 2.0 × 107 cells/mL, as displayed in Figure 4. The readsorption results were fit by the Hill equation, giving the maximum adsorptions for 5.0 and 7.0 wt % of 4 × 106 and 5.0 × 106 cells/cm2, respectively. The standard deviation in those adsorption measurements is approximately ±5%. A higher EVAL concentration (10.0 wt %) did not provide a higher recognition capacity (saturating at approximately 4.4 × 106 cells/cm2). For non-imprinted polymers (NIPs), the average maximum adsorption was 2.4 × 106 cells/cm2. In all cases (including NIPs), the Hill coefficient, which quantifies cooperativity in binding, was 2−3. The Hill equation is equivalent to the Langmuir equation when the Hill coefficient = 1, which would have indicated non-cooperative binding. It seems quite unlikely that conformational changes in the film lead to cooperativity in binding (especially with non-imprinted films); rather, the Hill coefficient likely reflects increasing algal aggregation in solution as the concentration is increased, leading to binding of increasing algal dimers, trimers, and larger aggregates.27 Interestingly, the highest EVAL concentration 14018
dx.doi.org/10.1021/la5031119 | Langmuir 2014, 30, 14014−14020
Langmuir
Article
Figure 5. (a) Time course measurements of the OCV of algae fuel cells using algae-imprinted EVAL-coated and bare Pt/ITO/PET electrodes as the anode, (b) OCV changes with different EVAL concentrations applied for AIP- and NIP-coated anodes in biofuel cells, (c) polarization behavior of the algal fuel cells, with voltage (open symbols) and power (filled symbols) changing with the loading of different resistances using AIP electrodes made with various polymer concentrations, and (d) maximum power related to EVAL concentrations used for microcontact imprinting.
■
The 7 wt % EVAL films gave the better adsorption than 5 wt % (Figure 4) and gave the best electrochemical performance, even though both films appeared to have nearly the same density of cavities. We hypothesize that the thinner film is unable to form cavities of the same quality or affinity, while thicker films offer no further improvement in binding and actually hinder H2 transport.
■
AUTHOR INFORMATION
Corresponding Author
*Telephone: +886-7-591-9455 and/or +886-912-178-751. Email: [email protected] and/or [email protected]. Notes
The authors declare no competing financial interest.
■
ACKNOWLEDGMENTS The authors acknowledge the National Science Council of the Republic of China (ROC), Taiwan, for financially supporting this research under Contracts NSC 102-2220-E-390-001 and NSC 102-2220-E-006-004 and the Ministry of Science and Technology of the ROC under Contracts MOST 103-2220-E390-001 and MOST 103-2220-E-006-007.
CONCLUSION
The present work reveals that algae-imprinted polymers made by 7.0 wt % EVAL that contains 38 mol % ethylene were better able to recognize algae than other polymer concentrations. In this specific polymer concentration, the average pore diameter of the algae-imprinted polymer was 10.69 ± 1.07 μm, to provide the highest saturated absorption of algae. An electrochemical performance comparison between the algaeimprinted EVAL-coated and bare Pt/ITO/PET electrodes indicated that the former achieved a power output of 1.65± 0.16 mW/m2 at a current density of 9.10 mA/m2; this power is substantially higher than that obtained with bare or NIP-coated algal electrodes. These results suggest that the polymer microenvironment may increase the metabolic activity of algae.
■
REFERENCES
(1) Ren, K.; Banaei, N.; Zare, R. N. Sorting inactivated cells using cell-imprinted polymer thin films. ACS Nano 2013, 7, 6031−6036. (2) Mahmoudi, M.; Bonakdar, S.; Shokrgozar, M. A.; Aghaverdi, H.; Hartmann, R.; Pick, A.; Witte, G.; Parak, W. J. Cell-imprinted substrates direct the fate of stem cells. ACS Nano 2013, 7, 8379−8384. (3) Chen, W.-J.; Lee, M.-H.; Thomas, J. L.; Lu, P.-H.; Li, M.-H.; Lin, H.-Y. Microcontact imprinting of algae on poly(ethylene-co-vinyl
14019
dx.doi.org/10.1021/la5031119 | Langmuir 2014, 30, 14014−14020
Langmuir
Article
alcohol) for biofuel cells. ACS Appl. Mater. Interfaces 2013, 5, 11123− 11128. (4) Borovicka, J.; Stoyanov, S. D.; Paunov, V. N. Shape recognition of microbial cells by colloidal cell imprints. Nanoscale 2013, 5, 8560− 8568. (5) Ren, K.; Zare, R. N. Chemical recognition in cell-imprinted polymers. ACS Nano 2012, 6, 4314−4318. (6) Jeon, H.; Kim, G. Effects of a cell-imprinted poly(dimethylsiloxane) surface on the cellular activities of MG63 osteoblast-like cells: Preparation of a patterned surface, surface characterization, and bone mineralization. Langmuir 2012, 28, 13423−13430. (7) Schirhagl, R.; Hall, E. W.; Fuereder, I.; Zare, R. N. Separation of bacteria with imprinted polymeric films. Analyst 2012, 137, 1495− 1499. (8) Dickert, F. L.; Hayden, O.; Halikias, K. P. Synthetic receptors as sensor coatings for molecules and living cells. Analyst 2001, 126, 766− 771. (9) Hayden, O.; Bindeus, R.; Haderspöck, C.; Mann, K.-J.; Wirl, B.; Dickert, F. L. Mass-sensitive detection of cells, viruses and enzymes with artificial receptors. Sens. Actuators, B 2003, 91, 316−319. (10) Hayden, O.; Lieberzeit, P. A.; Blaas, D.; Dickert, F. L. Artificial antibodies for bioanalyte detectionSensing viruses and proteins. Adv. Funct. Mater. 2006, 16, 1269−1278. (11) Jenik, M.; Schirhagl, R.; Schirk, C.; Hayden, O.; Lieberzeit, P.; Blaas, D.; Paul, G.; Dickert, F. L. Sensing picornaviruses using molecular imprinting techniques on a quartz crystal microbalance. Anal. Chem. 2009, 81, 5320−5326. (12) Eersels, K.; van Grinsven, B.; Ethirajan, A.; Timmermans, S.; Jiménez Monroy, K. L.; Bogie, J. F. J.; Punniyakoti, S.; Vandenryt, T.; Hendriks, J. J. A.; Cleij, T. J.; Daemen, M. J. A. P.; Somers, V.; De Ceuninck, W.; Wagner, P. Selective identification of macrophages and cancer cells based on thermal transport through surface-imprinted polymer layers. ACS Appl. Mater. Interfaces 2013, 5, 7258−7267. (13) Borovička, J.; Metheringham, W. J.; Madden, L. A.; Walton, C. D.; Stoyanov, S. D.; Paunov, V. N. Photothermal colloid antibodies for shape-selective recognition and killing of microorganisms. J. Am. Chem. Soc. 2013, 135, 5282−5285. (14) Cohen, T.; Starosvetsky, J.; Cheruti, U.; Armon, R. Whole cell imprinting in sol−gel thin films for bacterial recognition in liquids: Macromolecular fingerprinting. Int. J. Mol. Sci. 2010, 11, 1236−1252. (15) Dickert, F. L.; Hayden, O. Bioimprinting of polymers and sol− gel phases. Selective detection of yeasts with imprinted polymers. Anal. Chem. 2002, 74, 1302−1306. (16) Luo, Y.; Liu, G.; Zhang, R.; Zhang, C. Power generation from furfural using the microbial fuel cell. J. Power Sources 2010, 195, 190− 194. (17) Catal, T.; Li, K.; Bermek, H.; Liu, H. Electricity production from twelve monosaccharides using microbial fuel cells. J. Power Sources 2008, 175, 196−200. (18) Rezaei, F.; Xing, D.; Wagner, R.; Regan, J. M.; Richard, T. L.; Logan, B. E. Simultaneous cellulose degradation and electricity production by Enterobacter cloacae in a microbial fuel cell. Appl. Environ. Microbiol. 2009, 75, 3673−3678. (19) Wang, X.; Cheng, S.; Feng, Y.; Merrill, M. D.; Saito, T.; Logan, B. E. Use of carbon mesh anodes and the effect of different pretreatment methods on power production in microbial fuel cells. Environ. Sci. Technol. 2009, 43, 6870−6874. (20) Patil, S. A.; Surakasi, V. P.; Koul, S.; Ijmulwar, S.; Vivek, A.; Shouche, Y. S.; Kapadnis, B. P. Electricity generation using chocolate industry wastewater and its treatment in activated sludge based microbial fuel cell and analysis of developed microbial community in the anode chamber. Bioresour. Technol. 2009, 100, 5132−5139. (21) Dumas, C.; Mollica, A.; Féron, D.; Basséguy, R.; Etcheverry, L.; Bergel, A. Marine microbial fuel cell: Use of stainless steel electrodes as anode and cathode materials. Electrochim. Acta 2007, 53, 468−473. (22) Rosenbaum, M.; Schröder, U.; Scholz, F. Utilizing the green alga Chlamydomonas reinhardtii for microbial electricity generation: A living solar cell. Appl. Microbiol. Biotechnol. 2005, 68, 753−756.
(23) Lin, H.-Y.; Hsu, C.-Y.; Thomas, J. L.; Wang, S.-E.; Chen, H.-C.; Chou, T.-C. The microcontact imprinting of proteins: The effect of cross-linking monomers for lysozyme, ribonuclease A and myoglobin. Biosens. Bioelectron. 2006, 22, 534−543. (24) Ghirardi, M. L.; Zhang, L.; Lee, J. W.; Flynn, T.; Seibert, M.; Greenbaum, E.; Melis, A. Microalgae: A green source of renewable H2. Trends Biotechnol. 2000, 18, 506−511. (25) Niemann, J. Unraveling fuel cell electrical measurements. Fuel Cell Mag. 2005, April/May, 26−31. (26) Min, B.; Logan, B. E. Continuous electricity generation from domestic wastewater and organic substrates in a flat plate microbial fuel cell. Environ. Sci. Technol. 2004, 38, 5809−5814. (27) Ackleh, A. S.; Fitzpatrick, B. G. Modeling aggregation and growth processes in an algal population model: Analysis and computations. J. Math. Biol. 1997, 35, 480−502. (28) Thauer, R. K.; Jungermann, K.; Decker, K. Energy conservation in chemotrophic anaerobic bacteria. Bacteriol. Rev. 1977, 41, 100−180.
14020
dx.doi.org/10.1021/la5031119 | Langmuir 2014, 30, 14014−14020
