Redox-Cycling and H2O2 Generation by Fabricated Catecholic Films

Feb 14, 2011 - Foyer , C. H.; Noctor , G. Plant Cell 2005, 17, 1866– 1875 ... I.; Foster , S. J.; Asensi , A.; Taybi , T.; Barnes , J.; Foyer , C. H...
0 downloads 0 Views 5MB Size
Download PDF
ARTICLE pubs.acs.org/Biomac

Redox-Cycling and H2O2 Generation by Fabricated Catecholic Films in the Absence of Enzymes Eunkyoung Kim,† Yi Liu,† C. Jacyn Baker,|| Robert Owens,|| Shunyuan Xiao,†,‡ William E. Bentley,†,§ and Gregory F. Payne*,†,§ Center for Biosystems Research, ‡Plant Science and Landscape Architecture, and §Fischell Department of Bioengineering, University of Maryland, 5115 Plant Sciences Building, College Park, Maryland 20742, United States Beltsville Agricultural Research Center, United States Department of Agriculture, Beltsville, Maryland 20705, United States

)

†

bS Supporting Information ABSTRACT: Phenolic matrices are ubiquitous in nature (e.g., lignin, melanin, and humics) but remain largely intractable to characterize. We examined an abiotic phenol-polysaccharide matrix fabricated by the anodic grafting of catechol to chitosan films. Previous studies have shown that catechol-modified chitosan films are redox-active and can be repeatedly interconverted between oxidized and reduced states. Here we developed quantitative electrochemical methods to characterize biorelevant redox properties of the catechol-modified chitosan films. Our analysis demonstrates that these films can (i) accept electrons from biological reductants (e.g., ascorbate and nicotinamide adenine dinucleotide phosphate, NADPH) and (ii) donate electrons in a model biological oxidation process. Furthermore, these films can donate electrons to O2 to generate H2O2. The demonstration that abiotic catechol-chitosan films possess catalytic activities in the absence of enzymes suggests the possibility that phenolic matrices may play an important role in redox cycling and reactive oxygen species (ROS) signaling in biology and the environment.

’ INTRODUCTION Well-characterized electron transfer operations in biology involve either sequential steps among spatially organized proteins for energy harvesting (e.g., respiration and photosynthesis) or the transfer of reducing equivalents between distant proteins mediated by soluble cofactors (e.g., nicotinamide adenine dinucleotide phosphate, NADPH). Electron transfer operations that are less well-characterized also exist in nature and they include the oxidative burst during the innate immune response of plants and insects and the extracellular transfer of electrons in the environment. Interestingly, macromolecular phenolics are commonly reported to be coincident with these redox activities; lignin and melanin synthesis are often observed in plant1 and insect2,3 defense, and humics are often observed in redox-active soils.4 Phenolic matrices are ubiquitous in nature, but their insolubility, heterogeneity, and structural complexity make them difficult to study.5-8 As a result, seemingly simple questions remain unresolved (e.g., Is lignin a highly organized or random polymer?9 Or are melanin’s properties the result of large conjugated polymers or aggregates of lower molecular weight phenolics?10). Furthermore, the biological activities of phenolic matrices can be context-dependent (e.g., neuromelanins can be either pro- or antioxidants,11 and melanin at the host-pathogen interface can benefit the host2,3 or the pathogen12-14). Because of the difficulty in studying phenolic matrices in vivo and the r 2011 American Chemical Society

challenge of extracting biological matrices intact, it is common to investigate abiotically synthesized phenolic matrices to suggest putative biological activities.6,7,15,16 In previous studies, we fabricated an abiotic catecholpolysaccharide matrix using the method illustrated in Scheme 1a.17 The pH-responsive film-forming aminopolysaccharide chitosan18-21 is first electrodeposited from solution onto an electrode surface;22 then, the chitosan-coated electrode is anodically modified with catechol. During film modification, Scheme 1a illustrates that catechol diffuses through the chitosan film and is anodically oxidized at the electrode surface, and the oxidation product grafts to chitosan by reactions that include the formation of Michael’s-type adducts or Schiff bases (putative structure at the right after rearrangement).17,23-25 The photograph in Scheme 1b shows films prepared on gold-coated silicon wafers. Previous studies have shown that the catecholmodified chitosan films are nonconducting but redox-active. Specifically, the catechol-modified chitosan films cannot transfer electrons directly with the underlying electrode but can repeatedly donate and accept electrons with appropriate soluble mediators (ferrocene and ruthenium complexes).26 The ability of these films to exchange electrons with soluble Received: September 28, 2010 Revised: January 25, 2011 Published: February 14, 2011 880

dx.doi.org/10.1021/bm101499a | Biomacromolecules 2011, 12, 880–888

Biomacromolecules

ARTICLE

Scheme 1. Fabrication and Behavior of Catechol-Modified Chitosan Filmsa

Gold-coated substrates were prepared by sputtering 15 nm chromium, followed by 200 nm of gold on a 4 in diameter silicon wafer, as previously described.27 Before each experiment, these gold-coated electrodes were cleaned with piranha solution (30% H2O2/70% H2SO4), rinsed with water, and dried with nitrogen gas. (WARNING: Piranha reacts violently with organics.) Electrochemical Instrumentation. Chitosan was electrodeposited onto gold-coated electrodes using a DC power supply (model 6614C Agilent Technologies) with the gold-coated electrode serving as the cathode and a Pt foil serving as the anode in a two-electrode system. Electrochemical measurements (cyclic voltammetry and chronocoulometry) were performed using a three-electrode system with Ag/AgCl as a reference electrode and Pt wire as a counterelectrode (CHI Instruments 6273C electrochemical analyzer). Film Fabrication and Surface Characterization. Chitosan was electrodeposited onto the gold-coated electrode by immersing the electrode into a chitosan solution (1%, pH 5.6) and applying a cathodic voltage to achieve a constant current density (3 A/m2, 30 s) using a DC power supply. The chitosan-coated electrode was washed with water and phosphate buffer and dried. For electrochemical oxidation, the chitosancoated electrode was immersed in a catechol solution (5 mM unless otherwise noted), with gold serving as the working electrode (anodic voltage of þ0.6 V, 5 min) in a three-electrode system. Chitosan-coated electrodes were prepared in triplicate with differing extents of catechol modification using catechol solutions of 2, 5, 10, and 20 mM. Chitosan and catechol-modified chitosan films were prepared as described above and then freeze-dried by immersion in liquid nitrogen. The thickness of the dried films was measured using a profilometer (P20h surface profilometer, TENCOR Instruments). The surface morphology of the dried films was measured using scanning electron microscopy (SU-70 SEM, Hitachi) and atomic force microscopy (Multimode 3 Scanning Probe Microscope, Veeco) with a Si cantilever (300 kHz, 40 N/m) operated in the tapping mode. The mean roughness of films in AFM image is calculated using Nanoscope Software (5.3v).

a

(a) Fabrication starts with a chitosan-coated electrode and modification of the chitosan film is initiated by the anodic oxidation of catechol. (b) Photographs of the electrodes.

mediators but not with the electrode confers interesting functional properties to the films (e.g., for amplification, partial-rectification, and switching).26 Here we report interactions between the abiotic catecholmodified chitosan films and redox systems common to biology and the environment. Specifically, we developed an electrochemical method to “titrate” the film’s redox state and discovered that this catechol-polysaccharide matrix can exchange electrons with common biological oxidizing and biological reducing systems. Furthermore, these catechol-modified films possess context-dependent activities for redox cycling and reactive oxygen species (ROS) generation.

Experimental Procedure for Assessing H2O2-generating Ability of the Films. Details of the experimental procedure for assessing the H2O2-generating ability of the catechol-modified chitosan film are provided in the Supporting Information. In brief, films were fabricated, converted into the appropriate state (oxidized or reduced), rinsed, immediately dried in a stream of N2 gas, and stored in an N2-filled tube. For each experiment, a film was incubated in a cuvette containing 200 μL of H2O for 15 min, after which the liquid was removed and the H2O2 level was determined. In addition, for a catalase treatment, 20 U/ mL of catalase was added to 100 μL of the liquid obtained from the reduced film and incubated for 30 min at room temperature, after which the resulting liquid was removed and analyzed for H2O2. As indicated, experiments were performed using air, N2, or O2 gaseous environments.

’ RESULTS AND DISCUSSION

’ EXPERIMENTAL SECTION

Film Fabrication and Surface Characterization. We fabricated chitosan films to have differing extents of catechol modification using two steps. First, chitosan was electrodeposited onto the gold-coated electrode by immersing the electrode into a chitosan solution (1%, pH 5.6) and applying a cathodic voltage to achieve a constant current density (3 A/m2, 30 s). The chitosancoated electrode was washed with water and phosphate buffer and then dried. Second, the chitosan-coated electrode was immersed into a catechol solution, connected as the working electrode in a three-electrode system (Pt counter and Ag/AgCl reference), and biased to a constant anodic voltage of þ0.6 V for 5 min. During electrochemical catechol-modification, the anodic R charge transfer (q = i dt) was measured by chronocoulometry,

Materials. The water (>18 MΩ) used in this study was obtained from a Super Q water system (Millipore). Chitosan from crab shells (85% deacetylation and 200 kDa), catechol, KH2PO4, K2HPO4, Ru(NH3)6Cl3 (Ru3þ), 1,10 -ferrocenedimethanol (Fc), and acetosyringone (40 -hydroxy-30 50 -dimethoxyacetophenone) were purchased from Sigma-Aldrich. Chitosan solutions were prepared by dissolving chitosan flakes in HCl to achieve a final pH of 5 to 6, as previously described.27 All solutions of catechol and mediators (Fc and Ru3þ) were prepared in phosphate buffer (0.1 M; pH 7.0). Analysis of H2O2 was performed by the ferrous mediated oxidation in the xylenol orange reaction using standard assay kit (PeroXoquant Quantitative Peroxide Assay Kits, Pierce, IL). 881

dx.doi.org/10.1021/bm101499a |Biomacromolecules 2011, 12, 880–888

Biomacromolecules

ARTICLE

porosity of the chitosan film and is consistent with previous observations.17 Electrochemical Switching of the Film’s Redox State. In previous studies, we observed that the catechol-modified chitosan films could be converted to their oxidized or reduced states by brief electrochemical treatments that presumably convert the grafted moieties into catechols (reduced state designated QH2) or quinones (oxidized state designated Q).26 For oxidation to the Q state, the catechol-modified chitosan film was immersed in a solution of the mediator ferrocene dimethanol (Fc; 50 μM, 0.1 M phosphate buffer), and an anodic potential of þ0.5 V was applied to the underlying gold electrode for 2 min using a three-electrode system. (Chemical details of this oxidation are provided in the Electron-Accepting Properties of the Film section.) For reduction to the QH2 state, the catecholmodified chitosan film was immersed in a solution of the Ru(NH3)6Cl3 mediator (Ru3þ; 50 μM, 0.1 M phosphate buffer). and a cathodic potential of -0.4 V was applied for 2 min using the same three-electrode system. (Chemical details of this reduction are provided in the Electron-Donating Properties of the Film section.) After either treatment, the film-coated electrode was rinsed with water to remove the mediator. The above electrochemical method for switching the redox state of the films also allows a quantitative “titration” of the number of donatable electrons or the number of available electron-accepting sites.28 Specifically, the charge transfer during the 2 min of electrochemical switching was monitored by chronocoloumetry (see below). Electron-Accepting Properties of the Film. Electrochemical Methodology to Analyze Quantitatively the Films. Chronocoulometry under oxidizing conditions provides a method to “titrate” the number of electrons that can be “extracted” from the film. This analytical method is illustrated in Figure 2 and employs the Fc mediator and an anodic potential (þ0.5 V) for 2 min. The analytical negative control is a catechol-modified film that had been previously oxidized to its Q state, and the behavior of this negative control is schematically illustrated in Figure 2a. The Fc mediator diffuses from the bulk solution through the oxidized film, Fc undergoes anodic oxidation to Fcþ at the electrode, and the Fcþ product diffuses through the film and into the bulk solution. The small anodic charge transfer that is observed during this 2 min period for this negative control (qNC) is illustrated in Figure 2c and is attributed to diffusion of Fc (qDiff). The analytical positive control is a catechol-modified film that had been previously reduced to its QH2 state, and the behavior of this positive control is schematically illustrated in Figure 2b. In this case, the Fc mediator diffuses through the reduced film to be anodically oxidized to Fcþ; however, the Fcþ product can either diffuse back to the bulk or be rereduced (i.e., recycled) to Fc by accepting electrons from the film. Therefore, for this positive control, the observed charge transfer under these oxidizing conditions (qPC) includes contributions from both Fc diffusion (qDiff) and Fc recycling in the film (qFilm,max). Figure 2c illustrates that a considerably larger charge transfer is observed for this positive control. As illustrated in Figure 2c, the difference in charge transfer between the positive and negative controls (qFilm,max = qPC qNC) is a measure of the maximum number of donateable electrons that could be transferred between the film and the Fc mediator. (This analysis assumes that extraction of electrons from the film during this 2 min of chronocoulometric titration is complete.) For comparison, it is useful to convert the charge

Figure 1. Film fabrication. (a) Extent of catechol modification (NCatechol; mol/cm2) can be controlled by the concentration of catechol used during the anodic modification step. (b) SEM images of films of unmodified chitosan and catechol-modified chitosan (modification with 5 mM catechol at þ0.6 V for 5 min). (c) AFM images of films of unmodified chitosan and catechol-modified chitosan.

and the observed charge transfer (q; C) was converted into the extent of modification (NCatechol; mol/cm2) using the Faraday equation (eq 1) with the assumptions that (i) all anodic charge transfer is due to catechol oxidation, (ii) two electrons are transferred per molecule of catechol oxidized (n = 2), and (iii) each oxidized catechol grafts to chitosan. q ð1Þ N ¼ nAF In eq 1, A is the electrode area (cm2) and F is the Faraday constant (96 485 C/mol). Figure 1a indicates that NCatechol can be systematically varied by varying the catechol concentration used during electrochemical modification. Also, the relatively small error bars for the triplicate films in Figure 1a indicate that film preparation is reasonably reproducible. The morphology of dried films prepared from 5 mM catechol was examined using SEM and AFM. The SEM images in Figure 1b show a relatively smooth surface for the chitosan films, whereas the surface of the catechol-modified film is considerably rougher. The AFM images in Figure 1c further indicate that catechol modification increases surface roughness with estimated mean roughness values of 2 nm for the chitosan film and 11 nm for the catechol-modified chitosan film. Also, profilometry measurements indicate that the catechol-modified chitosan film is considerably thicker than the chitosan film (750 ( 30 vs 400 ( 20 nm). Together, these measurements suggest that a moderate extent of catechol-modification increases the volume and 882

dx.doi.org/10.1021/bm101499a |Biomacromolecules 2011, 12, 880–888

Biomacromolecules

ARTICLE

Figure 3. Correlation between the film’s redox capacity (NFilm,max) and number of grafted catechol moieties (NCatechol).

Figure 2. Chronocoulometric titration under oxidizing conditions using ferrocene dimethanol (Fc). (a) Scheme of film reaction in an analytical negative control. (b) Scheme of film’s recycling reaction in an analytical positive control. (c) Chronocoulometry method to determine the capacity of the film to donate electrons (qFilm,max = qPC - qNC).

Figure 4. Catechol-modified chitosan films accept electrons from biological reductants. (a) Schematic of experiment where oxidized films are incubated for 5 min with 5 mM of either NADPH or ascorbate. (b) Analysis of the film’s redox state by chronocoulometric titration under oxidizing conditions (50 μM Fc; þ0.5 V). Titration of an oxidized and a reduced film are the negative and positive analytical controls, respectively. (c) Quantity of electrons accepted by the catechol-modified chitosan film.

transfer to the number of moles of electrons transferred (NFilm,max; mole e-/cm2) using eq 1, where n = 1. Intrinsic Redox Capacity of the Film. Intuitively, the film’s capacity to accept electrons is expected to be linearly dependent on the number of grafted catechol moieties. (Unmodified chitosan is not redox-active29.) To test this expectation, chitosancoated electrodes with differing extents of catechol modification (Figure 1) were treated as illustrated in Figure 2c to allow the determination of qFilm,max and NFilm,max. Figure 3 shows that NFilm,max increases as a function of catechol modification (NCatechol), although the dependence is nonlinear. Possibly this nonlinear behavior is due to a decrease in probe permeability for the highly modified films, as previously observed.26 Alternatively, the highly modified films may be overoxidized with a loss of redox-active substituents. Interestingly, a comparison of the x and y scales in Figure 3 suggests that only about 5-10% of the grafted catechol moieties remain redox-active even for the lightly modified chitosan. Presumably, the complex grafting reactions result in some structures (e.g., phenolic oligomers) that lack the substituents required for reversible redox activity. Nevertheless, the results in Figure 3 indicate that the extent of catechol modification and the film’s redox capacity can be systematically controlled by experimental conditions.

Films Accept Electrons from Ascorbate and NADPH. We examined whether the catechol-modified chitosan films could accept electrons from the common biological reductants NADPH or ascorbate, as illustrated in Figure 4a. The experimental catechol-modified chitosan film (modified using 5 mM catechol) was initially converted to a fully oxidized (Q) state (like the analytical negative control), incubated with either NADPH (5 mM) or ascorbate (5 mM) for 5 min, rinsed, and then analyzed by electrochemical “titration” under oxidizing conditions. Figure 4b shows that the titration results for these experimental films are similar to that for the analytical positive control, indicating that the catechol-modified chitosan films had accepted electrons from NADPH or ascorbate, and subsequently donated these electrons during the analytical titration. Figure 4b also shows results from an experimental control in which an oxidized film was incubated for 15 min in phosphate buffer without reductant and then titrated by chronocoulometry. 883

dx.doi.org/10.1021/bm101499a |Biomacromolecules 2011, 12, 880–888

Biomacromolecules

ARTICLE

Figure 6. Catechol-modified chitosan films donate electrons during enzymatic redox cycling. (a) Experimental model of the biological redox cycling of acetosyringone (AS) with horseradish peroxidase (HRP). (b) Analysis of the film’s redox state by chronocoulometric titration under reducing conditions (50 μM Ru3þ; -0.4 V). Titration of a reduced and an oxidized film are the negative and positive analytical controls, respectively. (c) Semiquantitative analysis of electrons donated by the film.

Figure 5. Chronocoulometric titration under reducing conditions using Ru(NH3)63þ. (a) Scheme of film reaction in an analytical negative control. (b) Scheme of film’s recycling reaction in an analytical positive control. (c) Chronocoulometry graph to semiquantitatively analyze the film’s oxidation state.

the film’s oxidation state to be assessed. However, experimental constraints (background currents associated with O2 reduction) limit this approach to semiquantitative analysis useful for comparative purposes. As indicated in Figure 5, chronocoulometry under reducing conditions is performed using the Ru(NH3)63þ (Ru3þ) mediator and a cathodic potential (-0.4 V) for 2 min. The analytical negative control in this case is a catecholmodified film that was previously reduced to its QH2 state, and the behavior of this negative control is schematically illustrated in Figure 5a. The Ru3þ mediator diffuses from the bulk solution through the reduced film, Ru3þ undergoes cathodic reduction to Ru2þ at the electrode, and the Ru2þ product diffuses through the film and into the bulk solution. The “titration” curve in Figure 5c shows a relatively large cathodic charge transfer for this negative control (vs the negative control in Figure 2c), and this is attributed to the background cathodic currents. For semiquantitative analysis, we assign the observed charge transfer for this negative control (qNC) to the diffusion of the Ru3þ mediator (qDiff) plus a background charge transfer (qBackground), as illustrated in Figure 5c. The analytical positive control is a catechol-modified film that had been previously oxidized to its Q state, and the behavior of this positive control is schematically illustrated in Figure 5b. In this case, the Ru3þ mediator diffuses through the oxidized film to be cathodically reduced to Ru2þ, and this Ru2þ product can

The similarity of the titration between this buffer control and the analytical negative control indicates that no electrons were transferred to the film in the absence of reductant. Figure 4c shows the number of titratable electrons accepted by the catechol-modified film during the 5 min of treatment with NADPH or ascorbate (NFilm,accept). We calculated this value by converting charge transfer measured by chronocoulometry into a molar basis using eq 1 and subtracting the results from the negative control from those observed for the experimental film (NFilm,accept = NObsd - NNC). The results from this experiment demonstrate that the catechol-modified chitosan films can accept electrons from biological reductants, and this electron transfer occurs rapidly (during the 5 min incubation) in the absence of enzymes. Potential Relevance to Biology. The observations in Figure 4 are consistent with suggestions that the redox cycling of some quinone-based agents may deplete intracellular NAD(P)H.30 Furthermore, the rapid donation of electrons from ascorbate to our phenolic matrix may provide an explanation for the observation that ascorbate in the plant apoplast exists predominantly in an oxidized, dehydroascorbate form. (Lignin in the cell wall possibly accepts electrons from apoplastic ascorbate.)31-33 Electron-Donating Properties of the Film. Electrochemical Methodology to Semiquantitatively Analyze the Films. In principle, chronocoulometry under reducing conditions should allow 884

dx.doi.org/10.1021/bm101499a |Biomacromolecules 2011, 12, 880–888

Biomacromolecules

ARTICLE

(1 mM) in phosphate buffer (0.1 M; pH 7.0) for 5 min. After this enzymatic treatment, the film was analyzed by chronocoulometric titration under reducing conditions. Figure 6b shows that this “ASþHRPþH2O2” film was depleted of electrons similar to that of the positive control. This result indicates that most of the electrons had been donated from the film during the enzymatic redox cycling of AS. Finally, Figure 6b shows the titration results from four control films, which were treated with two of the three components of this system or simply with phosphate buffer. Similar values of cathodic charge transfer were observed for the titration of these controls, and these control values are considerably smaller than those for the experimental film. Figure 6c provides a semiquantitative summary of the net electron donation (relative to the negative control; NObs - NNC) by the catechol-modified chitosan films in this in vitro model oxidation system. Films Undergo Non-Enzymatic Redox Cycling. To demonstrate redox cycling in the absence of enzymes, we electrochemically oxidized AS by a film-coated electrode as illustrated in Figure 7a. The controls in this experiment are electrodes coated with either an unmodified chitosan film or a catechol-modified film after conversion to its oxidized (Q) state. The cyclic voltammograms (Figure 7b) and chronocoulometric measurements (Figure 7c) for these controls show relatively small oxidation currents and anodic charge transfer. The experimental electrode with the catechol-modified film converted into its reduced (QH2) state shows a considerable amplification of both the oxidative current and anodic charge transfer. This result indicates that the catechol-modified chitosan films can donate electrons to electrochemically oxidized AS and thus participate in nonenzymatic redox cycling. Potential Relevance to Biology. The direct relevance of the observed redox cycling to putative roles for AS in delignification and plant defense is impossible to assess. However, it is interesting to note that AS serves as a soluble phenolic mediator to “extract” electrons from the catechol-modified chitosan films. Recent studies also report that phenolics may serve as redox mediators for extracellular electron transport (e.g., for anaerobic respiration),40-45 and this may be important for bioremediation46-48 and microbial fuel cells.49-51 In addition, the observed redox cycling in the absence of enzymes is consistent with hypotheses put forth to explain the mode-of-action of some quinone-based anticancer agents52 and the deleterious effects of some air pollutants.53 H2O2 Generation by the Films. Films Donate Electrons to O2 to Generate H2O2. The final studies examined the possibility that the catechol-modified chitosan films could donate electrons to O2 to generate reactive oxygen species (ROS), as illustrated in Figure 8a. Electrodes with catechol-modified chitosan films were electrochemically converted to their reduced (QH2) state and then incubated in water at room temperature for 15 min, after which the solution was analyzed for H2O2 by the ferrous mediated oxidation in the xylenol orange method.54 Figure 8b shows that the solution H2O2 level was 3.7 ( 0.1 μM. It is interesting to note that this H2O2 value is similar to those reported for stressed plants.55 Treatment of the solution with catalase (20 U/ml for 30 min) depleted the H2O2, as indicated in Figure 8b. Two control films were also tested; one unmodified chitosan and a second catechol-modified chitosan that had been electrochemically oxidized (to Q state). Minimal H2O2 generation was observed with these control films.

Figure 7. (a) Schematic of abiotic redox cycling by anodic oxidation of AS (50 μM). (b) Cyclic voltammetry indicates an amplification of AS oxidation currents for a catechol-modified chitosan film in its reduced state (QH2). (c) Chronocoulometric measurements (at þ0.7 V) show increased anodic charge transfer for AS oxidation with a catecholmodified chitosan film in its reduced state (QH2).

either diffuse to the bulk or be reoxidized (i.e., recycled) to Ru3þ by donating electrons to the film. Therefore, for this positive control, the measured charge transfer under these reducing conditions (qPC) includes contributions from diffusion (qDiff), background (qBackground), and recycling within the film (qFilm). As expected, Figure 5c illustrates that a considerably larger charge transfer is observed for this positive control (compared with the negative control in Figure 5c). Films Donate Electrons in Bio-Relevant Oxidizing Systems. To demonstrate that the catechol-modified chitosan films can donate electrons in a biologically relevant oxidation processes, we created the experimental model illustrated in Figure 6a. Acetosyringone (AS), a low-molecular-weight derivative of lignin, can be enzymatically oxidized, and its oxidized form has been suggested to be a diffusible mediator for delignification in biological (e.g., fungal) and technological operations.34-37 Interestingly, a similar system may be involved in a plant innate immune response. When tobacco cells are elicited by bacterial pathogens, they generate extracellular AS,38 and this phenolic is subsequently consumed during an oxidative burst, presumably by peroxidase enzymes.39 Figure 6a illustrates that we examined the possibility that the catechol-modified film could donate electrons to the oxidized AS generated from a peroxidase reaction. The experimental film in Figure 6 was initially reduced to its QH2 state (like the negative control) and then incubated with AS (1 mM), horseradish peroxidase (HRP; 5 U/mL), and H2O2 885

dx.doi.org/10.1021/bm101499a |Biomacromolecules 2011, 12, 880–888

Biomacromolecules

ARTICLE

Figure 9. Correlation between electron-donation for H2O2-generation (NH2O2) and the film’s redox capacity (NFilm,max measured electrochemically under oxidizing conditions).

enlist phenolics (i.e., melanin) to inactivate the ROS generated by the host’s immune cells,12-14,57,58 whereas some host cells appear to employ phenolics to enhance pathogen damage by redox cycling.2,3 Whereas the uses of ROS for effector functions in immunity are well-established, there is emerging evidence that ROS (especially H2O2) may also perform signaling functions.31,32,59,60 The observation reported here that the abiotic catechol-chitosan matrix can generate H2O2 in the absence of enzymes raises interesting questions. Are the phenolic matrices that are generated as part of the plant and insect innate immune response capable of generating ROS as an extracellular effector mechanism (e.g., in the plant apoplast61 or the insect hemolymph2,3)? Also, if H2O2 is generated nonenzymatically, then can it also perform signaling functions? Correlation between Electron Donation for H2O2 Generation and the Film’s Redox Capacity. The results in Figure 3 indicate that reduced films can donate electrons to the soluble Fc mediator, whereas results in Figure 8 show that the reduced films can donate electrons to O2 to generate H2O2. Figure 9 shows a cross-correlation between these two electron-donating capabilities. In this correlation, the film’s ability to donate electrons to the electrochemical mediator is quantified by the redox capacity NFilm,max, as measured by chronocoulometric titration under oxidizing conditions (Figure 3). To facilitate comparison, the film’s ability to donate electrons to O2 to generate H2O2 is calculated using the following equation.

Figure 8. Catechol-modified chitosan films donate electrons for ROSgeneration. (a) Schematic of electron transfer from film to O2 for H2O2generation. (b) Observed H2O2 level for 1 cm2 films incubated in water (200 μL) for 15 min. (c) Observed H2O2 level when films were incubated in water (15 min) in contact with differing gases. (d) Observed H2O2 level when films with differing extents of catechol modification (NCatechol) were incubated with water (15 min in contact with air).

N H2 O2 ¼

Figure 8c shows results for films incubated for 15 min with solutions that were saturated with air, O2, or N2. As expected, H2O2 generation increased with an increasing O2 content of the gas. The ability of the films to donate electrons to O2 should also depend on the number of grafted catechol moieties. To test this possibility, we generated chitosan films with varying extents of catechol modification, electrochemically converted these films into either their oxidized (Q) or reduced (QH2) states, and incubated them for 15 min in water (in contact with air). After incubation, the H2O2 levels were determined, as shown in Figure 8d. As expected, H2O2 generation increased monotonically with the extent of catechol-modification (NCatechol) provided that the films were in their reduced form. Potential Relevance to Biology. The ability of catecholmodified chitosan matrices to donate electrons to O2 to generate H2O2 provides a mechanistic link between the redox properties of phenolics and the generation of ROS.56 Such a link is especially interesting in pathobiology because ROSs are commonly used to perform effector functions in hostpathogen interactions. Interestingly, some pathogens appear to

nCV A

ð2Þ

In eq 2, NH2O2 (mole e-/cm2) is the number of moles of electrons donated to generate H2O2, n is the number of electrons donated per H2O2 (n = 2), C is the concentration of H2O2 measured, V is the solution volume (200 μL) used for incubation, and A is the exposed area of the film during the incubation. As expected, Figure 9 shows a strong correlation between both electron-donating activities (correlation coefficient of 0.94). Quantitatively, the 10-fold difference in electrochemical electron-donation versus the electron-donation for H2O2-generation may be because H2O2 is neither an initial nor a stable product of electron donation to O2. Furthermore, the 15 min of incubation in water may not be sufficient for the complete donation of electrons from the film to O2. (See the Supporting Information.) Nevertheless, this correlation supports the conclusion that the intrinsic redox activity of the grafted catechols endows these films with the ability to generate ROS in the absence of enzymes. 886

dx.doi.org/10.1021/bm101499a |Biomacromolecules 2011, 12, 880–888

Biomacromolecules

ARTICLE

’ CONCLUSIONS Abiotic catechol-modified chitosan films were fabricated and observed to donate and accept electrons nonenzymatically with biologically relevant oxidants and reductants under ambient conditions. Also, these abiotic catechol-polysaccharide films can donate electrons in a context-dependent manner to generate ROS. The demonstration that these phenolic matrices possess catalytic activities in the absence of enzymes or carrier proteins could have important implications for host-pathogen interactions across lignin/melanin barriers2,3,12 and electron transfer in the environment and in biofuel cells.4,44,45,62 Furthermore, the observation of context-dependent redox activity and ROS-generation may be relevant to understanding the pro- versus antioxidant activities of neuromelanins,11,63 the mode-of-action of bactericidal antibiotics,64,65 and the possible nonenzymatic generation of ROS signaling molecules.66-68

(14) Nosanchuk, J. D.; Casadevall, A. Antimicrob. Agents Chemother. 2006, 50, 3519–3528. (15) Gidanian, S.; Farmer, P. J. J. Inorg. Biochem. 2002, 89, 54–60. (16) Toffoletti, A.; Conti, F.; Sandron, T.; Napolitano, A.; Panzella, L.; D’Ischia, M. Chem. Commun. 2009, 4977–4979. (17) Wu, L. Q.; Ghodssi, R.; Elabd, Y. A.; Payne, G. F. Adv. Funct. Mater. 2005, 15, 189–195. (18) Ladet, S.; David, L.; Domard, A. Nature 2008, 452, 76–79. (19) Redepenning, J.; Venkataraman, G.; Chen, J.; Stafford, N. J. Biomed. Mater. Res., Part A 2003, 66, 411–416. (20) Pang, X.; Zhitomirsky, I. Mater. Chem. Phys. 2005, 94, 245–251. (21) Luo, X. L.; Xu, J. J.; Du, Y.; Chen, H. Y. Anal. Biochem. 2004, 334, 284–289. (22) Yi, H.; Wu, L. Q.; Bentley, W. E.; Ghodssi, R.; Rubloff, G. W.; Culver, J. N.; Payne, G. F. Biomacromolecules 2005, 6, 2881–2894. (23) Kerwin, J. L.; Whitney, D. L.; Sheikh, A. Insect Biochem. Mol. Biol. 1999, 29, 599–607. (24) Wu, L. Q.; McDermott, M. K.; Zhu, C.; Ghodssi, R.; Payne, G. E. Adv. Funct. Mater. 2006, 16, 1967–1974. (25) Muzzarelli, C.; Muzzarelli, R. A. A. Trends Glycosci. Glycotechnol. 2002, 14, 223–229. (26) Kim, E.; Liu, Y.; Shi, X.-W.; Yang, X.; Bentley, W. E.; Payne, G. F. Adv. Funct. Mater. 2010, 20, 2683–2694. (27) Wu, L. Q.; Yi, H. M.; Li, S.; Rubloff, G. W.; Bentley, W. E.; Ghodssi, R.; Payne, G. F. Langmuir 2003, 19, 519–524. (28) Aeschbacher, M.; Sander, M.; Schwarzenbach, R. P. Environ. Sci. Technol. 2010, 44, 87–93. (29) Zangmeister, R. A.; Park, J. J.; Rubloff, G. W.; Tarlov, M. J. Electrochim. Acta 2006, 51, 5324–5333. (30) Dragan, M.; Dixon, S. J.; Jaworski, E.; Chan, T. S.; O’Brien, P. J.; Wilson, J. X. Brain Res. 2006, 1078, 9–18. (31) Foyer, C. H.; Noctor, G. Plant Cell 2005, 17, 1866–1875. (32) Foyer, C. H.; Noctor, G. Plant Cell Environ. 2005, 28, 1056– 1071. (33) Pignocchi, C.; Kiddle, G.; Hernandez, I.; Foster, S. J.; Asensi, A.; Taybi, T.; Barnes, J.; Foyer, C. H. Plant Physiol. 2006, 141, 423–435. (34) Camarero, S.; Ibarra, D.; Martinez, A. T.; Romero, J.; Gutierrez, A.; del Rio, J. C. Enzyme Microb. Technol. 2007, 40, 1264–1271. (35) Camarero, S.; Canas, A. I.; Nousiainen, P.; Record, E.; Lomascolo, A.; Martinez, M. J.; Martinez, A. T. Environ. Sci. Technol. 2008, 42, 6703–6709. (36) Camarero, S.; Ibarra, D.; Martinez, M. J.; Martinez, A. T. Appl. Environ. Microbiol. 2005, 71, 1775–1784. (37) Gutierrez, A.; Rencoret, J.; Ibarra, D.; Molina, S.; Camarero, S.; Romero, J.; Del Rio, J. C.; Martinez, A. T. Environ. Sci. Technol. 2007, 41, 4124–4129. (38) Baker, C. J.; Whitaker, B. D.; Roberts, D. P.; Mock, N. M.; Rice, C. P.; Deahl, K. L.; Aver’yanov, A. A. Physiol. Mol. Plant Pathol. 2005, 66, 90–98. (39) Baker, C. J.; Roberts, D. P.; Mock, N. M.; Whitaker, B. D.; Deahl, K. L.; Aver’yanov, A. A. Physiol. Mol. Plant Pathol. 2005, 67, 296–303. (40) Turick, C. E.; Beliaev, A. S.; Zakrajsek, B. A.; Reardon, C. L.; Lowy, D. A.; Poppy, T. E.; Maloney, A.; Ekechukwu, A. A. FEMS Microbiol. Ecol. 2009, 68, 223–235. (41) Turick, C. E.; Caccavo, F.; Tisa, L. S. FEMS Microbiol. Lett. 2003, 220, 99–104. (42) Turick, C. E.; Tisa, L. S.; Caccavo, F. Appl. Environ. Microbiol. 2002, 68, 2436–2444. (43) Stams, A. J. M.; de Bok, F. A. M.; Plugge, C. M.; van Eekert, M. H. A.; Dolfing, J.; Schraa, G. Environ. Microbiol. 2006, 8, 371–382. (44) Newman, D. K.; Kolter, R. Nature 2000, 405, 94–97. (45) Hatch, J. L.; Finneran, K. T. Curr. Microbiol. 2008, 56, 268–273. (46) O’Loughlin, E. J. Environ. Sci. Technol. 2008, 42, 6876–6882. (47) Van der Zee, F. R.; Cervantes, F. J. Biotechnol. Adv. 2009, 27, 256–277. (48) Perminova, I. V.; Kovalenko, A. N.; Schmitt-Kopplin, P.; Hatfield, K.; Hertkorn, N.; Belyaeva, E. Y.; Petrosyan, V. S. Environ. Sci. Technol. 2005, 39, 8518–8524.

’ ASSOCIATED CONTENT

bS

Supporting Information. Schematics of our experimental methods and systems, lengthy descriptions of our experimental procedures, and additional plots of our experimental results (with additional controls). This material is available free of charge via the Internet at http://pubs.acs.org.

’ AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected]. Tel: 301-405-8389. Fax: 301-3149075.

’ ACKNOWLEDGMENT We gratefully acknowledge financial support from the Robert W. Deutsch Foundation, the National Science Foundation (NSF; EFRI-0735987), the Department of Defense, Defense Threat Reduction Agency (W91B9480520121), and the Office of Naval Research (N000141010446). ’ REFERENCES (1) Huckelhoven, R. Annu. Rev. Phytopathol. 2007, 45, 101–127. (2) Nappi, A. J.; Christensen, B. M. Insect Biochem. Mol. Biol. 2005, 35, 443–459. (3) Christensen, B. M.; Li, J.; Chen, C. C.; Nappi, A. J. Trends Parasitol 2005, 21, 192–199. (4) Roden, E. E.; Kappler, A.; Bauer, I.; Jiang, J.; Paul, A.; Stoesser, R.; Konishi, H.; Xu, H. F. Nature Geoscience 2010, 3, 417–421. (5) Bothma, J. P.; de Boor, J.; Divakar, U.; Schwenn, P. E.; Meredith, P. Adv. Mater. 2008, 20, 3539–3542. (6) d’Ischia, M.; Napolitano, A.; Pezzella, A.; Meredith, P.; Sarna, T. Angew. Chem., Int. Ed. 2009, 48, 3914–3921. (7) Meredith, P.; Powell, B. J.; Riesz, J.; Nighswander-Rempel, S. P.; Pederson, M. R.; Moore, E. G. Soft Matter. 2006, 2, 37–44. (8) Meredith, P.; Sarna, T. Pigm. Cell Res. 2006, 19, 572–594. (9) Davin, L. B.; Lewis, N. G. Curr. Opin. Biotechnol. 2005, 16, 407– 415. (10) Pezzella, A.; Iadonisi, A.; Valerio, S.; Panzella, L.; Napolitano, A.; Adinolfi, M.; d’Ischia, M. J. Am. Chem. Soc. 2009, 131, 15270–15275. (11) Simon, J. D.; Hong, L.; Peles, D. N. J. Phys. Chem. B 2008, 112, 13201–13217. (12) Jacobson, E. S. Clin. Microbiol. Rev. 2000, 13, 708–717. (13) Jacobson, E. S.; Hong, J. D. J. Bacteriol. 1997, 179, 5340–5346. 887

dx.doi.org/10.1021/bm101499a |Biomacromolecules 2011, 12, 880–888

Biomacromolecules

ARTICLE

(49) Bond, D. R.; Lovley, D. R. Appl. Environ. Microbiol. 2005, 71, 2186–2189. (50) Lovley, D. R. Curr. Opin. Biotechnol. 2008, 19, 564–571. (51) Lovley, D. R. Geobiology 2008, 6, 225–231. (52) Gutierrez, P. L. Front. Biosci. 2000, 5, D629–D638. (53) Squadrito, G. L.; Cueto, R.; Dellinger, B.; Pryor, W. A. Free Radical Biol. Med. 2001, 31, 1132–1138. (54) Bou, R.; Codony, R.; Tres, A.; Decker, E. A.; Guardicila, F. Anal. Biochem. 2008, 377, 1–15. (55) Mittler, R. Trends Plant Sci. 2002, 7, 405–410. (56) Nakagawa, H.; Hasumi, K.; Woo, J. T.; Nagai, K.; Wachi, M. Carcinogenesis 2004, 25, 1567–1574. (57) Missall, T. A.; Lodge, J. K.; McEwen, J. E. Eukaryotic Cell 2004, 3, 835–846. (58) Chatfield, C. H.; Cianciotto, N. P. Infect. Immun. 2007, 75, 4062–4070. (59) Torres, M. A.; Jones, J. D. G.; Dangl, J. L. Plant Physiol. 2006, 141, 373–378. (60) Tschopp, J.; Schroder, K. Nat. Rev. Immunol 2010, 10, 210–215. (61) Bolwell, G. P.; Bindschedler, L. V.; Blee, K. A.; Butt, V. S.; Davies, D. R.; Gardner, S. L.; Gerrish, C.; Minibayeva, F. J. Exp. Bot. 2002, 53, 1367–1376. (62) Dietrich, L. E.; Teal, T. K.; Price-Whelan, A.; Newman, D. K. Science 2008, 321, 1203–1206. (63) Rao, K. S. J.; Hegde, M. L.; Anitha, S.; Musicco, M.; Zucca, F. A.; Turro, N. J.; Zecca, L. Prog. Neurobiol. 2006, 78, 364–373. (64) Kohanski, M. A.; Dwyer, D. J.; Collins, J. J. Nat. Rev. Microbiol. 2010, 8, 423–435. (65) Kohanski, M. A.; Dwyer, D. J.; Hayete, B.; Lawrence, C. A.; Collins, J. J. Cell 2007, 130, 797–810. (66) Paulsen, C. E.; Carroll, K. S. ACS Chem. Biol. 2009, 5, 47–62. (67) Van Breusegem, F.; Dat, J. F. Plant Physiol. 2006, 141, 384–390. (68) Miller, G.; Schlauch, K.; Tam, R.; Cortes, D.; Torres, M. A.; Shulaev, V.; Dangl, J. L.; Mittler, R. Sci. Signaling 2009, 2, A26–A35.

888

dx.doi.org/10.1021/bm101499a |Biomacromolecules 2011, 12, 880–888