pubs.acs.org/Langmuir © 2009 American Chemical Society
Graphene Oxide-Facilitated Electron Transfer of Metalloproteins at Electrode Surfaces Xiaolei Zuo, Shijiang He, Di Li, Cheng Peng, Qing Huang, Shiping Song, and Chunhai Fan* Laboratory of Physical Biology, Shanghai Institute of Applied Physics, Chinese Academy of Sciences, Shanghai 201800, China Received July 10, 2009. Revised Manuscript Received August 4, 2009 Graphene is a particularly useful nanomaterial that has shown great promise in nanoelectronics. Because of the ultrahigh electron mobility of graphene and its unique surface properties such as one-atom thickness and irreversible protein adsorption at surfaces, graphene-based materials might serve as an ideal platform for accommodating proteins and facilitating protein electron transfer. In this work, we demonstrate that graphene oxide (GO) supports the efficient electrical wiring the redox centers of several heme-containing metalloproteins (cytochrome c, myoglobin, and horseradish peroxidase) to the electrode. Importantly, proteins retain their structural intactness and biological activity upon forming mixtures with GO. These important features imply the promising applications of GO/protein complexes in the development of biosensors and biofuel cells.
Introduction 1,2
Graphene is a monolayer of tightly packed carbon atoms. This two-dimensional (2D) carbon nanomaterial has become an excitingly new material in physical sciences due to its excellent electronic, thermal, and mechanical features.3-5 Particularly, graphene has found important applications in nanoelectronics due to the fast mobility of electrons (120 000 cm2/(V s)).6-8 However, the “chemistry part” of graphene and its oxidation form, graphene oxide (GO),9 as well as their biological applications remain to be extensively explored.10,11 In this work, we interrogate the effect of GO on the electron transfer of several heme-containing metalloproteins at electrode surfaces. Metalloproteins are well-known to participate in a number of important biological processes, e.g., photosynthesis and bioenergetic metabolism.12 During the past several decades, it has become a popular topic to study metalloproteins adsorbed at electrode surfaces, which mimics protein-protein binding and proteinmediated electron transfer (ET) occurring in physiological conditions.13 Also importantly, surface-confined metalloproteins have found widespread applications in biosensors and bioelectronics.14,15 Along these lines, great effort has been taken to
design functionalized surfaces that allow efficient in vitro ET between redox centers of proteins and surfaces. While redox centers of proteins are often buried within folded polypeptide shells, which are usually regarded as poor electron conductors, a number of metalloproteins have been found to undergo direct ET at various surfaces since the seminal work of Hill and co-workers.16 Earlier effort was taken to either find appropriate electrode surfaces (e.g., edge-plane pyrolytic graphite or boron-doped diamond)17 or tailor metal electrode surface with appropriate functional groups (e.g., gold modified with selfassembled monolayers).18,19 More recently, nanomaterials-facilitated protein ET at electrode surfaces has attracted particular interest due to the availability of various nanoscale materials with excellent electrical conductivity and tunable morphology.19 For example, several groups have reported that both gold nanoparticles and carbon nanotubes can serve as electron relays to electrically wire a range of redox proteins (e.g., cytochrome c (cyt c),20-22 glucose oxidase (GOD),23,24 glutathione reductase,25 and hydrogenase26,27) to electrodes. Analogously, “molecular-wire”-like conjugated oligomers were also employed to wire the deeply buried redox site of a copper amine oxidase to a gold electrode.28
*Corresponding author. E-mail:
[email protected]. (1) Geim, A. K.; Novoselov, K. S. Nat. Mater. 2007, 6, 183–191. (2) Berger, C.; Song, Z. M.; Li, T. B.; Li, X. B.; Ogbazghi, A. Y.; Feng, R.; Dai, Z. T.; Marchenkov, A. N.; Conrad, E. H.; First, P. N.; de Heer, W. A. J. Phys. Chem. B 2004, 108, 19912–19916. (3) Li, D.; Kaner, R. B. Science 2008, 320, 1170–1171. (4) Westervelt, R. M. Science 2008, 320, 324–325. (5) Freitag, M. Nat. Nanotechnol. 2008, 3, 455–457. (6) Bolotin, K. I.; Sikes, K. J.; Hone, J.; Stormer, H. L.; Kim, P. Phys. Rev. Lett. 2008, 101, 096802. (7) Bolotin, K. I.; Sikes, K. J.; Jiang, Z.; Klima, M.; Fudenberg, G.; Hone, J.; Kim, P.; Stormer, H. L. Solid State Commun. 2008, 146, 351–355. (8) Tan, Y. W.; Zhang, Y.; Bolotin, K.; Zhao, Y.; Adam, S.; Hwang, E. H.; Das Sarma, S.; Stormer, H. L.; Kim, P. Phys. Rev. Lett. 2007, 99, 246803. (9) Park, S.; Ruoff, R. S. Nat. Nanotechnol. 2009, 4, 217–224. (10) Ruoff, R. Nat. Nanotechnol. 2008, 3, 10–11. (11) Sun, X.; Liu, Z.; Welsher, K.; Robinson, J. T.; Goodwin, A.; Zaric, S. Nano Res. 2008, 1, 203–212. (12) Voet, D.; Voet, J. G. Biochemistry, 2nd ed.; John Wiley & Sons: New York, 1993. (13) Jeuken, L. J. C. Biochim. Biophys. Acta 2003, 1604, 67–76. (14) Murphy, L. Curr. Opin. Chem. Biol. 2006, 10, 177–184. (15) Cracknell, J. A.; Vincent, K. A.; Armstrong, F. A. Chem. Rev. 2008, 108, 2439–2461.
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(16) Eddowes, M. J.; Hill, H. A. O. J. Chem. Soc., Chem. Commun. 1977, 771– 772. (17) Geng, R.; Zhao, G.; Liu, M.; Li, M. Biomaterials 2008, 29, 2794–2801. (18) Leger, C.; Bertrand, P. Chem. Rev. 2008, 108, 2379–2438. (19) Wang, C.; Yang, C.; Song, Y.; Gao, W.; Xia, X. Adv. Funct. Mater. 2005, 15, 1267–1275. (20) Brown, K. R.; Fox, A. P.; Natan, M. J. J. Am. Chem. Soc. 1996, 118, 1154– 1157. (21) Liu, H.; Tian, Y.; Deng, Z. Langmuir 2007, 23, 9487–9494. (22) Wang, J.; Li, M.; Shi, Z.; Li, N.; Gu, Z. Anal. Chem. 2002, 74, 1993–1997. (23) Xiao, Y.; Patolsky, F.; Katz, E.; Hainfeld, J. F.; Willner, I. Science 2003, 299, 1877–1881. (24) Sun, Y. Y.; Yan, F.; Yang, W. S.; Sun, C. Q. Biomaterials 2006, 27, 4042– 4049. (25) Scott, D.; Toney, M.; Muzikar, M. J. Am. Chem. Soc. 2008, 130, 865–874. (26) Alonso-Lomillo, M. A.; Rudiger, O.; Maroto-Valiente, A.; Velez, M.; Rodriguez-Ramos, I.; Munoz, F. J.; Fernandez, V. M.; De Lacey, A. L. Nano Lett. 2007, 7, 1603–1608. (27) McDonald, T. J.; Svedruzic, D.; Kim, Y. H.; Blackburn, J. L.; Zhang, S. B.; King, P. W.; Heben, M. J. Nano Lett. 2007, 7, 3528–3534. (28) Contakes, S. M.; Juda, G. A.; Langley, D. B.; Halpern-Manners, N. W.; Duff, A. P.; Dunn, A. R.; Gray, H. B.; Dooley, D. M.; Guss, J. M.; Freeman, H. C. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 13451–13456.
Published on Web 08/20/2009
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Figure 1. Cartoon for the GO-supported heme proteins at the surface of GC electrodes (oxygenated functional groups at the surface of GO are not shown for simplicity).
Armstrong and co-workers reported that microsized graphite particles can accommodate a range of redox proteins and allow rapid ET through the graphite layer.29 Aspired by their finding, we reason that GO might be a promising material for accommodating proteins and facilitating protein ET due to its unique surface properties such as one-atom thickness1,2 and irreversible protein adsorption ability due to strong hydrophobic interaction (like in the case of carbon nanotubes30-32), leading to hybrid nanobiomaterials that are useful in biosensors and biofuel cells. More recently, it was found that an ionic liquid modified graphene supported the direct electron transfer of GOD.33 In this work, we interrogated the effects of GO on electron transfer properties of several heme-containing metalloproteins (or heme proteins) at electrode surfaces, including cyt c, myoglobin (Mb), and horseradish peroxidase (HRP) (Figure 1).
Materials and Methods Materials. HRP (EC 1.11.1.7), cyt c (from equine heart), and Mb (from equine heart) were obtained from Sigma. The stock solutions of these proteins were in 25 mM phosphate buffer (pH 7.4) and stored at 4 °C. All solutions were prepared with Nanopure water (18 MΩ cm-1) from a Millipore Milli-Q system. GO was synthesized from graphite powder according to a modified Hummers and Offeman method.34,35 Briefly, graphite powder (4 g) was oxidized by adding to a hot solution (80 °C) of concentrated H2SO4 (24 mL) containing K2S2O8 (8 g) and P2O5 (8 g). The resultant dark blue mixture was thermally isolated and slowly cooled to room temperature over a period of 6 h. The mixture was diluted to 300 mL and then filtrated with a 0.22 μm filter membrane (Generay Biotech Co., Ltd., Shanghai, China). The filtered product was dried overnight at 60 °C. The preoxidized graphite powder (2 g) was added to 92 mL of cold H2SO4 (0 °C), and 12 g of KMnO4 was gradually added under stirring in an ice bath. After being stirred for 15 min, 2 g of NaNO3 was added to (29) Vincent, K. A.; Li, X.; Blanford, C. F.; Belsey, N. A.; Weiner, J. H.; Armstrong, F. A. Nat. Chem. Biol. 2007, 3, 761–762. (30) Balavoine, F.; Schultz, P.; Richard, C.; Mallouh, V.; Ebbesen, T. W.; Mioskowski, C. Angew. Chem., Int. Ed. 1999, 38, 1912–1915. (31) Chen, R. J.; Choi, H. C.; Bangsaruntip, S.; Yenilmez, E.; Tang, X. W.; Wang, Q.; Chang, Y. L.; Dai, H. J. J. Am. Chem. Soc. 2004, 126, 1563–1568. (32) Azamian, B. R.; Davis, J. J.; Coleman, K. S.; Bagshaw, C. B.; Green, M. L. H. J. Am. Chem. Soc. 2002, 124, 12664–12665. (33) Shan, C.; Yang, H.; Song, J.; Han, D.; Ivaska, A.; Niu, L. Anal. Chem. 2009, 81, 2378–2382. (34) Hummers, W.; Offeman, J. R. J. Am. Chem. Soc. 1958, 80, 1339–1339. (35) Kovtyukhova, N. I.; Ollivier, P. J.; Martin, B. R.; Mallouk, T. E.; Chizhik, S. A.; Buzaneva, E. V.; Gorchinskiy, A. D. Chem. Mater. 1999, 11, 771–778.
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Figure 2. Tapping-mode AFM image of as-prepared GO and the height profile along the dashed line in the panel. the mixture, and the resulting mixture was further stirred at 35 °C for 2 h and distilled water (200 mL) was added. The reaction was stopped with the addition of a mixture of 560 mL of distilled water and 10 mL of 30% H2O2. For purification, the mixture was washed with 1:10 HCl and then with water. The GO product was resuspended in water to a brown dispersion, which was subjected to dialysis to remove residual metal ions and acids. The purified GO dispersion was sonicated for 1.5 h at 300 W to exfoliate GO, and unexfoliated GO was removed by centrifugation (3000 rpm, 5 min). The as-prepared GO samples were then characterized with tapping-mode atomic force microscope (AFM)11,36 and transmission electron microscopy.37 Electrochemical Measurements. All electrochemical measurements were conducted at a CH Instruments (CHI 630). A conventional three-electrode cell, consisting of a modified glassy carbon (GC) disk electrode working (2 mm in diameter, CH Instruments Inc.), a Pt coil auxiliary electrode, and an Ag/ AgCl (saturated with KCl) reference electrode, was used for electrochemical measurements. Phosphate buffer (25 mM, pH 7.4) was used as the electrolyte in all experiments and extensively purged with high-purity nitrogen before experiments. Working electrodes were consecutively polished by 1, 0.3, and 0.05 μm of alumina polishing compounds, rinsed with excess water and ethanol, and briefly sonicated prior to each experiment. Proteins (cyt c, HRP, or Mb) of 2 mg/mL and GO (0.3 mg/mL) were mixed for 5 min at room temperature. Then 6 μL of the mixed solution was drop-cast on electrodes which were allowed to be dried at room temperature.
Results and Discussion GO, the oxide form of graphene, was employed in this work as it is water-dispersible and appropriate for biological applications. GO was synthesized from graphite powder as described in the literature, which is graphene functionalized with hydroxyl and carboxylic groups. The thickness of as-prepared GO is of ∼1 nm as measured by AFM, characteristic of a fully exfoliated graphene oxide sheet (Figure 2).38 We also find that the size of GO is fairly polydispersed with lateral dimension ranging from nanometers to micrometers. (36) Li, D.; Muller, M. B.; Gilje, S.; Kaner, R. B.; Wallace, G. G. Nat. Nanotechnol. 2008, 3, 101–105. (37) Li, X.; Zhang, G.; Bai, X.; Sun, X.; Wang, X.; Wang, E.; Dai, H. Nat. Nanotechnol. 2008, 3, 538–542. (38) Xu, Y.; Bai, H.; Lu, G.; Li, C.; Shi, G. J. Am. Chem. Soc. 2008, 130, 5856– 5857.
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Figure 3. (a) Cyclic voltammograms for the GO/cyt c modified electrode at various scan rates. Scan rates: 20, 30, 40, 50, 70, and 100 mV/s (from inner to outside curves). Inset: the anodic peak current as a function of scan rate. (b) Cyclic voltammograms for the GO/cyt c modified electrode obtained in the absence (solid line) and presence of 10 μM H2O2 (dotted line); scan rate: 50 mV/s. (c) Cyclic voltammograms for cyt c at a bare GC electrode.
Figure 4. (a) UV-vis spectra for native cyt c (solid line) and cyt c in the presence of GO (dashed line). (b) Fluorescence spectra for native cyt c (solid line) and cyt c in the presence of GO (dashed line).
Cyt c is probably the most studied protein in electrochemistry.39 While the direct, unmediated electrochemistry has been obtained at the surface of various functionalized electrodes and at specific bare electrode surfaces (e.g., edge-plane pyrolytic graphite), we find that GC does not support direct ET of cyt c since no reduction and oxidation peaks are observed in voltammograms using GC electrodes (Figure 3c). Interestingly, when cyt c is mixed with GO and adsorbed at the GC surface, a pair of well-defined CV peaks appears, corresponding to the reduction-oxidation conversion between the Fe(II) and the Fe(III) states of heme of cyt c (Figure 3a). Of note, GO alone does not produce any observable CV features in the potential window of interest. These peaks are stable, with minimal change over hundreds of potential scans. This high stability may arise from both strong hydrophobic and electrostatic interactions between cyt c and GO. The peak separation between the reduction and the oxidation peaks is relatively large (∼68 mV even at a slow scan rate of 20 mV/s). This phenomenon is often observed in heterogeneous protein film electrodes, possibly arising from the existence of thermodynamic and/or kinetic dispersion at the electrode surface.40 Peak currents are found to be linearly proportional to scan rates, suggesting a surface-confined electrode process (inset of Figure 3a).40 The apparent standard potential (the midpoint potential of reduction and oxidation peak potentials) is calculated to be 65 mV vs Ag/AgCl (264 mV vs NHE), which coincides well with the redox potential of native cyt c in solution.41 The electron (39) Beissenhirtz, M. K.; Scheller, F. W.; Stocklein, W. F.; Kurth, D. G.; Mohwald, H.; Lisdat, F. Angew. Chem., Int. Ed. 2004, 43, 4357–4360. (40) Nassar, A. E. F.; Zhang, Z.; Yu, N.; Rusling, J. F.; Kumosinski, T. F. J. Phys. Chem. B 1997, 101, 2224–2231. (41) Haymond, S.; Babcock, G. T.; Swain, G. M. J. Am. Chem. Soc. 2002, 124, 10634–10635. (42) Fan, C.; Gillespie, B.; Wang, G.; Heeger, A. J.; Plaxco, K. W. J. Phys. Chem. B 2002, 106, 11375–11383.
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transfer rate of cyt c is estimated to be 12.6 s-1 based on the reported method.42 It is important to examine conformational variation of cyt c upon interaction with GO. Absorption spectra provide information for different conformational states of cyt c. UV-vis spectra show that both the Soret band (408 nm) and the Q-band (∼529 nm) which are characteristic of native cyt c remain unaltered in the presence of GO (Figure 4a).42 Fluorescence studies also confirm the intactness of cyt c. We only observe small, background tryptophan fluorescence (Figure 4b), which is in direct contrast to unfolded cyt c produces a strong emission at ∼350 nm. This suggests Trp59 remains proximal to the heme group of cyt c in both the absence and presence of GO, leading to efficient fluorescence quenching via energy transfer.42 Previous studies have proven that native cyt c does not contain any peroxidase activity; however, very small structural variation of cyt c that cannot be readily probed in spectroscopy can lead to significant increase the peroxidase activity of cyt c toward the catalytic reduction of hydrogen peroxide (H2O2).43 Thus, the peroxidase activity of cyt c provides an extremely sensitive probe for the structural intactness of cyt c.43,44 We then test the GC electrode with cyt c and GO in a solution of H2O2 and find that the CVs are not significantly changed even at a high concentration of 100 μM H2O2 (without any electrocatalytic feature), suggesting that the GO-bound cyt c retains its intact structure even under this rigorous interrogation (Figure 3b). It is worthwhile to note that while a range of nanomaterials have been reported to facilitate the electrochemistry of cyt c, significant peroxidase activity of cyt c was observed with many of these materials (43) Diederix, R. E. M.; Ubbink, M.; Canters, G. W. ChemBioChem 2002, 1, 110–112. (44) Diederix, R. E. M.; Ubbink, M.; Canters, G. W. Biochemistry 2002, 41, 13067–13077.
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Figure 5. (a) Cyclic voltammograms for GO/Mb modified electrodes modified electrodes. Scan rate: 5 mV/s. (b) Cyclic voltammograms for GO/Mb modified electrode in the absence and presence of 1, 5, 10, 15, 20, 30, and 40 μM of H2O2 (from inner to outside curves). Inset: plot of the electrocatalytic current versus H2O2 concentration for GO/Mb modified electrodes.
Figure 6. (a) Cyclic voltammograms for GO/HRP modified electrodes. Scan rate: 50 mV/s. (b) Amperometric curves for GO/HRP modified electrodes with different concentrations of H2O2: (a) 0, (b) 1, (c) 5, (d) 10, (e) 15, (f) 20, and (g) 25 μM. Inset: plot of the current versus H2O2 concentration for GO/HRP modified electrodes.
(e.g., gold nanoparticles,21 carbon nanotubes,45 and mesoporous niobium oxide46), suggesting significant perturbation of the cyt c structure. We next interrogate the effects of GO on an oxygen storage protein, Mb. Mb is similarly mixed with GO and deposited at the surface of GC electrodes. Mb itself does not show any electrochemical activity at GC electrodes. However, we observe a pair of well-defined redox peaks for both Mb and HRP in the presence of GO, with the midpoint potential of -112 mV (vs Ag/AgCl). While Mb is not an enzyme, it possesses intrinsic peroxidase activity (in contrast to cyt c). Coincidently, we find that the GO/ Mb modified GC electrodes exhibit significant catalytic activity toward H2O2 (Figure 5), as characterized by an apparent increase in the reduction peak of Mb. This Mb-based sensor responds to H2O2 in the dynamic range of 1-40 μM. The electron transfer of a natural peroxidase, HRP, can also be enhanced with the aid of GO. As shown in Figure 6, we observe a pair of HRP reduction and oxidation peaks in the presence GO, with a midpoint potential at -70 mV (vs Ag/AgCl). HRP retains its peroxidase activity upon binding with GO, and this catalytic property provides a means to sensitively detect H2O2. (45) Zhao, G. C.; Yin, Z. Z.; Wei, X. W. Electrochem. Commun. 2005, 7, 256– 260. (46) Xu, X.; Tian, B.; Kong, J.; Zhang, S.; Liu, B.; Zhao, D. Adv. Mater. 2003, 15, 1932–1936.
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We employed amperometry to directly measure the current associated with the HRP-catalyzed electrochemical process. The potential is held at the catalytic reduction potential for H2O2 (-80 mV vs Ag/AgCl); a decay curve for current (I) vs time (t) is observed instantly after the onset of the potential, which rapidly reaches a plateau (steady-state current) within ∼20 s (Figure 6b). The reduction current of H2O2 is found to gradually increase along with the H2O2 concentration in the range of 1-25 μM. We have demonstrated that GO supports the efficient electrical wiring the redox centers of proteins to the electrode. In our experiments, GO effectively facilitates the ET of several heme proteins, leading to well-defined electrochemistry at GC electrodes. Importantly, proteins retain their structural intactness and biological activity upon forming mixtures with GO. These important features suggest that GO might be a promising nanomaterial in a range of areas involving metalloproteins, such as biosensors, bioelectronics, and biofuel cells. Acknowledgment. This work supported by the National Natural Science Foundation (20873175 and 20725516), Ministry of Science and Technology (2006CB933000, 2007CB936000, 2007AA06A406), Shanghai Municipal Commission for Science and Technology (0752 nm021, 0952 nm04600), and Ministry of Health (2009ZX10004-301).
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