Electrochemical Characterization of Pore Formation by Bacterial

Apr 11, 2008 - The interaction of pore-forming streptolysin O (SLO) with biomimetic lipid membranes has been studied by electrochemical methods...
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Langmuir 2008, 24, 5615-5621

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Electrochemical Characterization of Pore Formation by Bacterial Protein Toxins on Hybrid Supported Membranes Thomas Wilkop, Danke Xu,† and Quan Cheng* Department of Chemistry, UniVersity of California, RiVerside, California 92521 ReceiVed December 25, 2007. ReVised Manuscript ReceiVed February 21, 2008 The interaction of pore-forming streptolysin O (SLO) with biomimetic lipid membranes has been studied by electrochemical methods. Phosphatidylcholine lipid vesicles were deposited onto gold electrodes modified with supporting layers of hexyl thioctate (HT) or thioctic acid tri(ethylene glycol) ester (TA-TEGE), and integrity and permeability of the resulting membranes were characterized by cyclic voltammetry and impedance spectroscopy. Both positively and negatively charged electrochemical probes, potassium ferrocyanide, hexaammineruthenium(III) chloride, and ferrocene carboxylic acid (FCA), were employed to evaluate their suitability to probe the membrane permeability properties, with FCA exhibiting ideal behavior and thus employed throughout the work. Fusion of vesicles incubated with SLO on the electrodes yielded membranes that showed a distinctive response pattern for FCA as a function of SLO concentration. A direct dependence of both the currents and peak separation of FCA in the cyclic voltammograms was observed over a concentration range of 0–10 hemolytic units (HU)/µL of the toxin. The interaction of SLO with preformed supported lipid membranes was also investigated, and much lower response was observed, suggesting a different extent of membrane-toxin interactions on such an interface. Nonionic surfactant Triton was found to disrupt the vesicle structure but could not completely remove a preformed membrane to fully restore the electrode response. The information reported here offers some unique insight into toxin-surface interactions on a hybrid membrane, facilitating the development of electrochemically based sensing platforms for detecting trace amounts of bacterial toxins via the perforation process.

Introduction Membrane damaging toxins (MDTs) are some of the most prevalent toxins found in Nature. At least 35% of 325 protein toxins produced by Gram-negative and Gram-positive bacteria are membrane damaging toxins.1 Aside from the macroscopic effect of tissue and organ damage, many of the toxins contribute strongly to the virulence of bacteria. They play an important role in the bacterial pathogenesis in both humans and animals2–4 subverting host cell functions and altering the organism’s homeostasis and signal transduction pathways. The cumulative effect is an overall weakening of the organism’s innate defense mechanism, which in turn favors the multiplication of bacteria.5 Cell membrane disruption can also lead to the spreading of the bacteria through the tissue. MDTs can be grouped into three main classes:6 (a) toxins that enzymatically hydrolyze the phospholipids of the bilayer membrane of cells, (b) surfactantlike toxins that solubilize lipid membranes, and (c) pore-forming toxins. These toxins have been investigated with a variety of techniques, including artificial biomimetic membrane systems.7–10 Many pore forming toxins share a general working scheme: The released toxins bind to the specific receptor sites on the cell * Corresponding author, [email protected]. † Current address: Department of Biochemistry, Beijing Institute of Radiation Medicine, Beijing 100850, China. (1) Alouf, J. E. Pore-forming bacterial protein toxins: An overview. In Poreforming toxins; Current Topics in Microbiology and Immunology 257; Springer: Berlin, 2001; pp 1–14. (2) Paton, J. C. Trends Microbiol. 1996, 4, 103–106. (3) Titball, R. W. In Membrane damaging and cytotoxic phospholiases. The comprehensiVe sourcebook of bacterial protein toxins; Alouf, J. E., Freer, J. H., Eds.; Academic: London, 1999; pp 311-329. (4) Salyers, A. A.; Whitt, D. D.; Salyers, A. A.; Whitt, D. D. Bacterial pathogenesis: A molecular approach. In Bacterial pathogenesis: A molecular approach; ASM Press: Washington, DC, 1994. (5) Finlay, B. B.; Falkow, S. Microbiol. Mol. Biol. ReV. 1997, 61, 136. (6) Thelestam, M.; Mollby, R. Biochim. Biophys. Acta 1979, 557, 156–169. (7) Billington, S. J.; Jost, B. H.; Songer, J. G. FEMS Microbiol. Lett. 2000, 182, 197–205.

membrane and form noncovalently bound oligomeric structures (prepores). Upon some triggering mechanism, a structural change of the oligomeric complex occurs and the prepore inserts itself into the lipid membrane.8,11 The number of toxin monomers in the pore varies between 7 and 50.12 The size of the pores stretches over a large spectrum, from 1 to 2 nm in diameter for S. aureus R-toxin,13 Aeromonas aerolysin,14 and Vibrio El Tor hemolysin15 to 25-39 nm for streptolysin O8 and pneumolysin.16 The pores are nonselective and behave like molecular sieves, allowing only the egress of small particles and molecules out of the cell. The largest family of the pore forming toxins are the cholesterolbinding cytolysins. They are produced by at least 23 bacterial species from the genera Streptococcus, Bacillus, BreVibacillus, Paenibacillus, Clostridium, Listeria, and Arcanobacterium. These toxins have shown cytolytic activity for many eukaryotic cell types and share two common features: (a) the dependence of their cytolytic activity on the presents of cholesterol in the membrane and (b) the formation of large pores. (8) Menestrina, G.; Vecsey Semjen, B. In Biophysical methods and model membranes for the study of bacterial pore-forming toxins. The comprehensiVe sourcebook of bacterial protein toxins; Alouf, J. E., Freer, J. H., Eds.; Academic: London, 1999; pp 287-309. (9) Alouf, J. E. Pharm. Therapeu. 1980, 11, 661–717. (10) Sekiya, K.; Danbara, H.; Yase, K.; Futaesaku, Y. J. Bacteriol. 1996, 178, 6998–7002. (11) Lesieru, C.; Vecsey-Semjen, B.; Abrami, L.; Fivaz, M.; Van Der Goot, F. G. Mol. Membr. Biol. 1997, 14, 45–64. (12) Goot, F. G. v. d. In Bacterial Protein Toxins; Burns, D. L., Barbieri, J. T., Iglewski, B. H., Rappuoli, R., Eds.; ASM Press: Washington, DC, 2003; p 191. (13) Bhakdi, S.; Tranum-Jensen, J. Microbiol. ReV. 1991, 55, 733–750. (14) van der Goot, F. G. Trends Microbiol. 2000, 8, 89–90. (15) Shinoda, S. In Haemolysins of Vibrio cholerae and other Vibrio species. The comprehensiVe sourcebook of bacterial protein toxins; Alouf, J. E., Freer, J. H., Eds.; Academic: London, 1999; pp 373-385. (16) Gilbert, R. J. C.; Rossjohn, J.; Parker, M. W.; Tweten, R. K.; Morgan, P. J.; Mitchell, T. J.; Errington, N.; Rowe, A. J.; Andrew, P. W.; Byron, O. J. Mol. Biol. 1998, 284, 1223–1237.

10.1021/la704027c CCC: $40.75  2008 American Chemical Society Published on Web 04/11/2008

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We selected a member of this class, streptolysin O, as our model system to study the membrane perforation and its potential as a biosensing interface. Streptolysine O is a 69 kD β-hemolysin produced by the Gram-positive bacterium Streptococcus pyogenes.17 The bacterium Streptococcus pyogenes is the causative agent for a large number of prevalent diseases, such as acute pharyngitis, impetigo, rheumatic fever, and streptococcal toxic shock syndrome.18 In addition to their pathogenic significance, pore forming toxins have also received considerable attention for their utility in biotech19–23 and medical applications.24,25 Recent efforts have focused on the engineering of pore forming proteins for their application in biotherapeutics.26 In order to characterize the native activity of pore forming toxins, a biomimetic lipid membrane system is required. For the convenient interrogation of the membrane and toxin interaction by surface plasmon resonance,27 total internal reflection fluorescence microscopy,28 acoustic waveguides,29 or electrochemical techniques,30–33 it is essential that the membrane is formed directly on a substrate. Silicate surfaces facilitate the spontaneous fusion of vesicles into ideal bilayers.34 Unfortunately, the insulating nature of the glassy surfaces makes them unsuitable for electrochemical measurements. To allow the formation of lipid layers on gold substrates, surface modification is required, as vesicles do not fuse readily on unmodified gold surfaces. It has been shown that on alkanethiols with a hydrophilic headgroup, vesicle fusion results in bilayer membranes35 while hybrid membranes36 are formed on alkanethiols with hydrophobic headgroups. However, the inherently dense packing of these SAM layers prohibits an effective charge transfer37 and severely constrains their application for electrochemical measurements. To obtain a suitable support layer that allows electrochemical characterization of the membrane permeability, we have synthesized two disulfide-terminated compounds, hexyl thioctate (HT) and thioctic acid tri(ethylene glycol) ester (TA-TEGE), for surface modification. In this work, parameters that determine the behavior of the biomimetic lipid membranes on HT and TA(17) Bhakdi, S.; Roth, M.; Sziegoleit, A.; Tranumjensen, J. Infect. Immun. 1984, 46, 394–400. (18) Areschoug, T.; Carlsson, F.; Stalhammar-Carlemalm, M.; Lindahl, G. Vaccine 2004, 22, S9–S14. (19) Song, L.; Hobaugh, M. R.; Shustak, C.; Cheley, S.; Bayley, H.; Gouaux, J. E Science 1996, 274, 1859–1866. (20) Akeson, M.; Branton, D.; Kasianowicz, J. J.; Brandin, E.; Deamer, D. W. Biophys. J. 1999, 77, 3227–3233. (21) Krasilnikov, O. V.; Sabirov, R. Z.; Ternovsky, V. I.; Merzliak, P. G.; Tashmukhamedov, B. A. Gen. Physiol. Biophys. 1988, 7, 467–474. (22) Toby, A.; Jenkins, A.; Olds, J. A. Chem. Commun. 2004, 2106–2107. (23) Xu, D.; Cheng, Q. J. Am. Chem. Soc. 2002, 124, 14314–14315. (24) Giles, R. V.; Grzybowski, J.; Spiller, D. G.; Tidd, D. M. Nucleosides Nucleotides 1997, 16, 1155–1163. (25) Panchal, R. G.; Cusack, E.; Cheley, S.; Bayley, H. Nat. Biotechnol. 1996, 14, 852–856. (26) Panchal, R. G.; Smart, M. L.; Bowser, D. N.; Williams, D. A.; Petrou, S. Curr. Pharm. Biotechnol. 2002, 3, 99–115. (27) Rossi, C.; Homand, J.; Bauche, C.; Hamdi, H.; Ladant, D.; Chopineau, J. Biochemistry 2003, 42, 15273–15283. (28) Moran-Mirabal, J. M.; Edel, J. B.; Meyer, G. D.; Throckmorton, D.; Singh, A. K.; Craighead, H. G. Biophys. J. 2005, 89, 296–305. (29) Melzak, K. A.; Ellar, D. J.; Gizeli, E. Langmuir 2004, 20, 1386–1392. (30) Becucci, L.; Leon, R. R.; Moncelli, M. R.; Rovero, P.; Guidelli, R. Langmuir 2006, 22, 6644–6650. (31) Becucci, L.; Moncelli, M. R.; Guidelli, R. Langmuir 2006, 22, 1341– 1346. (32) Liu, X. H.; Huang, W. M.; Wang, E. K. J. Electroanal. Chem. 2005, 577, 349–354. (33) Becucci, L.; Guidelli, R.; Peggion, C.; Toniolo, C.; Moncelli, M. R. J. Electroanal. Chem. 2005, 576, 121–128. (34) Phillips, K. S.; Wilkop, T.; Wu, J. J.; Al-Kaysi, R. O.; Cheng, Q. J. Am. Chem. Soc. 2006, 128, 9590–9591. (35) Silin, V. I.; Wieder, H.; Woodward, J. T.; Valincius, G.; Offenhausser, A.; Plant, A. L. J. Am. Chem. Soc. 2002, 124, 14676–14683. (36) Plant, A. L.; Gueguetchkeri, M.; Yap, W. Biophys. J. 1994, 67, 1126– 1133. (37) Wang, Z. Z.; Wilkop, T.; Cheng, Q. Langmuir 2005, 21, 10292–10296.

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TEGE and their interactions with pore forming bacterial toxins are investigated, and the membrane integrity and its permeability are characterized by cyclic voltammetry (CV). Initial effort is placed on optimization of experimental conditions that lead to high-quality lipid membranes allowing for efficient electron transfer with the redox probes. We then evaluate the analytical performance of the surface to register the perforation of lipid membranes by SLO toxin. In addition to pore forming signals, we also investigate the influence of nonspecific protein interactions and the effect of surfactant on membrane integrity and response pattern, in an attempt to provide useful information toward further development of membrane-based protein assays.

Experimental Section Materials. L-R-Phosphatidylcholine (egg PC) was purchased from Avanti Polar Lipids (Alabaster, AL). Cholesterol, potassium ferrocyanide (K4Fe(CN)6), ferrocene carboxylic acid (FCA), hexaammineruthenium(III) chloride (Ru(NH3)6Cl3), dicyclohexyl carbodiimide, 4-dimethylaminopyridine, and 1-hexanol were from Aldrich. Anhydrous tri(ethylene glycol) was from Fluka. Streptolysin O (from Streptococcus pyogenes), bovine serum albumin, and Trizma were obtained from Sigma. All solutions were prepared with Milli-Q water (>18 MΩ). Synthesis of Hexyl Thioctate (HT) and Thioctic Acid Tri(ethylene glycol) Ester (TA-TEGE). To a stirred solution of thioctic acid (2.0 mmol) in 10 mL of methylene chloride, hexyl alcohol (2.0 mmol) (for the preparation of HT) or tri(ethylene glycol) anhydrous (10.0 mmol) (for the preparation of TA-TEGE) was added, respectively. The resulting mixture was stirred for 15 min at 0 °C (ice/water bath) under N2. Then dicyclohexyl carbodiimide (DCC, 4 mmol) and 4-(dimethylamino)pyridine (DMAP, 0.6 mmol) in cold CH2Cl2 (10 mL) were added to the above solution, and the mixture was allowed to react for 15 min at 0 °C, followed by 60 h reaction at room temperature. The resulting solution was filtered through a fine glass frit, and the clear filtrate was washed with water. The organic layer was separated and dried with MgSO4, filtered, and evaporated. The raw yellow oily product was further purified with silica gel (230–400 mesh, Sigma) and column chromatography using a mixture of CHCl3/methanol (97.5/2.5) as eluent. Collection, solvent removal and drying in vacuum yielded HT or TA-TEGE as yellow oils. NMR spectral data are as follows: 1H NMR (300 MHz, CDCl3). HT: δ 0.73–0.97 (t, 3H); 0.97–1.37 (m, 6H); 1.38–1.79 (m, 8H); 1.79–2.00 (m,1H); 2.16–2.34 (t, 2H); 2.34–2.54 (m, 1H); 2.96–3.28 (m, 2H); 3.43–3.64 (m, 1H); 3.90–4.15 (t, 2H). TA-TEGE: δ 1.37–1.89 (m, 6H); 1.82–2.11 (m, 3H); 2.39–2.48 (t, 2H); 2.49–2.56 (m, 1H); 3.00–3.29 (m, 3H); 3.44–3.82 (m, 8H); 4.09–4.35 (t, 2H). Preparation of Vesicles and Lipid Layer Modified Electrodes. Vesicles used for surface modification were prepared by probe sonication (model 250 sonifier, Baranson Ultrasonics). Egg PC and cholesterol suspended in chloroform were mixed in an amber vial in a 1:1 molar ratio. The organic solvent was then removed in a N2 stream to form a thin lipid layer. Appropriate amounts of 10 mM Tris buffer (pH 7.5 containing 150 mM NaCl) were then added, to bring the final lipid concentration to 1.0 mg/mL. The solution was sonicated at low amplitude at 0 °C for 30 min. After sonication, the resulting solution was filtered and stored at 4 °C for 1 h before use. The vesicles were always used within 36 h after preparation. The gold electrodes were prepared by thermal evaporation of a thin film of Au (ca. 200 nm) onto microscope glass slides with Cr as the adhesion layer. Prior to modification, the Au substrates were cleaned with piranha solution (Caution!), extensively rinsed in deionized (DI) water and dried in a N2 stream. The electrodes were then immersed in a 1.0 mM solution of either HT or TA-TEGE in ethanol for 2 h to form a self-assembled monolayer (SAM) and rinsed with ethanol and DI water. The contact angle measurements were conducted with a home-built device as described in a previous publication.38 (38) Phillips, K. S.; Cheng, Q. Anal. Chem. 2005, 77, 327–334.

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Two approaches were used to investigate the impact of the SLO toxin interaction on the electrochemical response of the lipid membrane. In both cases the SLO was first activated by incubation with 5 mM dithiothreitol for 1 h.39 In the first approach SLO was incubated with intact vesicles before the vesicles were fused. The 30 min incubation was carried out at room temperature, and then the mixture was transferred into the electrochemical cell for fusion on the SAM modified electrode. After another 30 min of incubation, the cell was rinsed twice with DI water and then with Tris buffer. In the second approach, vesicles were first fused on the SAM modified electrode, followed by addition of SLO and incubation for 30 min. The electrochemical cell was rinsed with DI water and buffer and filled with 150 µL of 1 mM FCA solution in Tris buffer for characterization. Extreme care was taken throughout the rinsing and measuring process to avoid the drying of the cell. Electrochemical Measurements. The electrochemical measurements were carried out with a three-electrode system using a CHI 650 electrochemical work station (CH Instruments, Austin, TX). The modified Au electrode formed part of the electrochemical cell and served as the working electrode. The active electrode area was defined by an O-ring (2 mm i.d.). An Ag/AgCl reference electrode and a platinum counter electrode completed the electrochemical system. Electron Microscopy. Transmission electron micrographs of the vesicles were obtained with a Philips CM 300 electron spectroscopic microscope. The image contrast was enhanced by staining the sample with 2% uranyl acetate. Pore formation by streptolysin O was assessed after incubating the vesicles for 30 min with 200 HU/µL SLO.

Results and Discussion 1. Electrochemical Behavior of Three Redox Probes on the SAM Modified Electrodes. The insertion of SLO toxin into lipid membranes was studied with two approaches: (a) by incubating vesicles in solution with the toxin and subsequently fusing the vesicles on the modified electrode and (b) by incubating an existing supported lipid membrane on the electrode with the toxin. Figure 1 shows a cartoon illustration of the experimental approaches and the chemical structure of TA-TEGE and HT used for electrode modification. In order to maximize the response in the membrane permeability study, two parameters of the membrane/probe system were optimized. One is the permeability of the supporting layer for the redox probe. Ideally the electron transfer kinetics on the supporting layer remains close to that of an untreated electrode. The other is the integrity of the pristine lipid membrane. Therefore, a defect-free surface coverage that provides an insulating layer on the electrode is desirable. The electron transfer process through the hybrid membrane system is an intricate interplay of the probes, supporting SAM and the lipid membrane. This led us to evaluate six discreet systems using three redox probes (potassium ferrocyanide, hexaammineruthenium(III) chloride, and ferrocene carboxylic acid) on two supporting surfaces (TA-TEGE and HT). The properties of the supporting layers have been found to strongly influence the spontaneous vesicle fusion and, therefore, determine the type and integrity of the lipid membrane formed on them.35 The effect of TA-TEGE and HT on surface hydrophobicity was first investigated. Adsorption of the ring tailgroups leads to a loosely packed SAM, yielding a highly pervious thin film for solvent and electrolytes. Their different chain lengths and degree of oxygenation should result in a different degree of hydrophobicity, which will strongly influence the initiation of spontaneous vesicle fusion. The higher degree of oxygenation in TA-TEGE should give the molecule a more polar characteristic and render it hydrophilic. This is confirmed by the experimental data. Sessile drop contact angle measurements determined angles of 60 ( 7° (39) Bernheimer, A. W.; Avigad, L. S. Infect. Immun. 1970, 1, 509–510.

Figure 1. (a) Structure of TA-TEGE and HT. (b) A cartoon illustration of the experimental approach.

for the untreated gold electrode, 51 ( 5° for the TA-TEGE SAM on Au, and 83 ( 6° for the HT SAM on Au. The highly negatively charged K4Fe(CN)6 probe was first used to characterize the electrochemical behavior on the membrane surface. Cyclic voltammograms were recorded for the supporting layer of TA-TEGE (Figure 2a) and HT (Figure 2b). On the bare Au electrode the electron transfer is relatively fast, as indicated by a peak separation of 114 mV. The formation of the TA-TEGE SAM clearly slowed down the kinetics, with the peak separation increasing to 445 mV. Both anodic and cathodic peak currents were reduced by half, and injection of vesicle solution on this surface further reduced the currents and increased the peak separation to 697 mV. Although the probe’s kinetics was slowed down, the membrane system still allowed the electrochemical reaction to proceed. Incomplete electrode coverage and/or a leaky membrane structure could account for this behavior. In comparison, the deposition of the HT support layer slows down the electron transfer more effectively. The cathodic peak was eliminated from the scanning window, and the magnitude of the anodic peak current was reduced slightly but shifted from +208 to +445 mV, indicating that the oxidation of Fe(CN)64experienced a substantial barrier. Vesicle fusion completely eliminated the anodic current, suggesting that the lipid layer blocked the electrode surface very effectively. The redox reaction of the positively charged Ru(NH3)6Cl3 is fast on the bare gold electrode (∆Ep ) 69 mV, Figure 3a).

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Figure 2. Cyclic voltammograms of 1.0 mM K4Fe(CN)6 on modified gold electrodes. The Au surface (solid line) was modified with TATEGE (a, dashed line) or HT (b, dashed line). The resulting surfaces were further modified by vesicle fusion (dotted lines). The electrolyte solution contained 10 mM Tris buffer (pH 7.5) and 150 mM NaCl. The scan rate was 0.1 V/s.

Electrode modification with TA-TEGE only marginally affected the response, with the peak separation remaining virtually identical and the peak current reduced only slightly. It is remarkable that after injection of vesicle solution the electrokinetics of the probe was hardly affected, and the peak currents decreased insignificantly (∼7%). The results again indicate incomplete surface coverage, likely due to poor vesicle fusion and/or instability of the resulting membrane. An interesting behavior was observed in the case of the HT electrode modification characterized with Ru(NH3)6Cl3. Despite the hydrophobic nature of the HT layer, the electrode kinetics of the probe was unchanged and the current magnitudes were reduced only by ca. 5%. This contrasts the behavior of straightchain alkanethiols such as octadecanthiol that have only one thiol terminus.36 The result demonstrates a high permeability of the HT supporting layer. In contrast to the TA-TEGE surface, injection of vesicle solution onto the HT electrode slowed down the kinetics. The anodic peak moved beyond 0.1 V and the cathodic peak shifted from –200 to –360 mV, while the current magnitudes decreased by 45% as compared to the HT surface (Figure 3b). From these results, Ru(NH3)6Cl3 appears to be only partially blocked on the supported membrane and is hence unsuitable for the intended permeability study.

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Figure 3. Cyclic voltammograms of 1.0 mM Ru(NH3)6Cl3 on modified gold electrodes. The Au surface (solid line) was first modified with TA-TEGE (a, dashed line) or HT (b, dashed line). The resulting surfaces were further modified by vesicle fusion (dotted lines). The electrolyte solution contained 10 mM Tris buffer (pH 7.5) and 150 mM NaCl. The scan rate was 0.1 V/s.

Ferrocene carboxylic acid is a small, negatively charged facile redox probe. On TA-TEGE modified surface, the peak current magnitude decreased, indicating that the electrode became less accessible to the probe (Figure 4a). The electrode kinetics, however, was unaffected with the peak separation remaining unchanged at 65 mV. Injection of vesicle solution reduced both anodic and cathodic peak currents by ca. 21% as compared to that of the TA-TEGE electrode. This behavior was similar to that observed for the Fe(CN)64- and Ru(NH3)6Cl3 probes. Taken together, we conclude that the hydrophilic TA-TEGE does not sustain a stable and complete lipid layer and, therefore, appears inadequate for membrane permeability studies. On the HT modified Au electrode, however, FCA essentially retains its fast kinetics, as is evident from the peak separation of 67 mV on the bare and 71 mV on the HT electrode (Figure 4b). A 13% reduction in anodic and cathodic peak currents was observed on HT. Vesicle fusion on the HT surface eliminated both anodic and cathodic current peaks, indicating that a well insulating lipid membrane against FCA has formed (Figure 4b). This behavior makes them the ideal system selected for membrane permeation assessments and thus were used throughout the investigation.

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Figure 5. Cyclic voltammetric responses for the supported membranes prepared by fusion of vesicles incubated with various concentrations of SLO toxin (9, blank; O, 1 HU/µL; 2, 2 HU/µL; 4, 4 HU/µL; (, 8 HU/µL; g, 10 HU/µL). The probe was 1.0 mM FCA in 10 mM Tris buffer (pH 7.5) and 150 mM NaCl. Scan rate was 0.1 V/s.

Figure 4. Cyclic voltammograms of 1.0 mM FCA on modified gold electrodes. The Au surface (solid line) was first modified with TATEGE (a, dashed line) and HT (b, dashed line), respectively. The resulting surfaces were further modified by vesicle fusion (dotted lines). The scan rate was 0.1 V/s.

2. The Effect of Pore Formation on the Electrochemical Response. The native behavior of the SLO toxin is the insertion of the protein molecules into the cholesterol-containing cell membrane of eucariotic cells. In an analytical context this condition is most closely emulated by the incubation of lipid vesicles with SLO in buffer. It is challenging to assess the magnitude of membrane permeation in suspended vesicles. In order to obtain a measurable signal, the incubated vesicles were fused on the HT modified electrode surface. Figure 5 shows the cyclic voltammograms obtained with 1 mM FCA as probe on various membrane surfaces. These membranes were formed through fusion of vesicles that were incubated with SLO concentrations from 0 to 10 HU/µL. HU is the hemolytic unit for SLO toxin and is defined as the amount of protein that lyses 50% of a 2% red blood cell suspension in phosphate-buffered saline at pH 7.4. Compared to the reference membrane formed from vesicles only, incubation with 1 HU/µL SLO dramatically increased the anodic current. At a potential of 550 mV the anodic current increased from 0.7 µA for the pristine lipid membrane to 2.38 µA. The position of the anodic peak at this high potential indicates a sluggish electrode kinetics. The cathodic current remained low, with the reduction peak outside the scanning window. For toxin concentrations of 2 and 4 HU/µL, an increase in both anodic and cathodic currents was observed. The anodic

peak positions shifted to lower potentials of 470 and 440 mV, respectively. Also the cathodic peak reappeared for toxin concentrations larger than 2 HU/µL, with a peak separation of 245 mV observed for 4 HU/µL SLO. For SLO concentrations of 8 and 10 HU/µL the electrode kinetics accelerated even further, though the current magnitudes no longer increased. This behavior suggests a diffusion layer overlap at high toxin concentrations,37 causing a saturation effect. The peak separation for 10 HU/µL SLO was 85 mV, close to that on the HT electrode (73 mV), while the redox current magnitudes were only about 10% smaller as compared to the HT surface. It appears that the membrane formed from incubation with 10 HU/µL has now obtained a degree of permeation that gives the probe a largely unrestricted access to the electrode. We believe that a certain degree of lateral diffusion beyond the pore diameter occurs, possibly due to the crown shape of the protruding part of the pores.40 The quantitative results from cyclic voltammetric analysis, including data not shown in Figure 5, are summarized in Table 1. The table highlights the fact that both peak separation and anodic current magnitudes are nonlinearly related to the toxin concentration, likely due to the “large” dynamic range studied. At higher concentration the peak separation offers a unique angle to estimate the toxin concentration, whereas at low hemolytic units the anodic current magnitude appears to be more suitable. In order to verify that the observed response is in fact caused by the formation of transmembrane pores and is not an artifact of incomplete vesicle fragments depositing, we characterized the vesicle structure with transmission electron microscopy. Representative micrographs of native vesicles are shown in Figure 6. The vesicles were of predominantly spherical shape, with varying diameters. The largest vesicles measured ca. 180 nm in diameter. Vesicles incubated with SLO toxin (Figure 6) clearly show the insertion of the toxin into the membrane and the formation of transmembrane pores. A variety of shapes including arcs and round pores were also observed. The fully developed pores, as can be well seen in Figure 6d, were large, with diameters of up to ca. 22 nm. The direct observation of the pores in the TEM micrographs supports the notion that the observed cyclic voltammetric response was due to the native pore forming activity of the SLO. (40) Sekiya, K.; Satoh, R.; Danbara, H.; Futaesaku, Y. J. Bacteriol. 1993, 175, 5953–5961.

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Table 1. Cyclic Voltammetric Results on the HT Modified Electrodes Fused with Vesicles That Have Been Incubated with Different Concentrations of SLO Toxin SLO 0 HU/µL Ep,a (mV)a Ip (µA)b ∆Ep (mV)c a

Anodic peak potential.

b

SLO 1.0 HU/µL

SLO 2.0 HU/µL

SLO 3.0 HU/µL

SLO 4.0 HU/µL

SLO 8.0 HU/µL

SLO 10.0 HU/µL

552 2.38

462 2.57

434 2.78 320

424 3.06 245

361 3.02 143

347 2.97 87

Peak current. c Peak separation.

Figure 7. Faradic impedance spectra for a lipid membrane before (squares) and after incubation (circles) with 1 HU/µL SLO. The impedance spectra were quantitatively analyzed using the equivalent circuit shown in the inset. The solid symbols refer to the real part of the impedance Z (left side axis) while the phase shift (open symbols) is shown in regard to the axis on the right side. The potential of the working electrode was +300 mV and the ac amplitude 5 mV. Figure 6. Transmission electron micrographs of pristine (top) and SLO incubated vesicles (bottom). Vesicles were incubated with 200 HU/µL SLO toxins. The scale bar measures 500 nm in top images and 60 nm in bottom images.

In the above study, perforated vesicles were fused on a HT supporting layer. An alternative approach, with the added benefit of allowing in situ analysis, is the induction of pore formation by SLO in a preformed supported membrane. For the pore formation process, the random movement of the membranebound SLO monomers seems to be important for the oligomerization process, which generates the prepores that eventually become the transmembrane pores. This requirement of membrane fluidity makes the post insertion into an existing membrane on a SAM layer a challenging task. We thus investigated if the HT support layer is suitable for such an approach. Cyclic voltammograms obtained with FCA for a pristine membrane and the identical membrane after incubation with 1 HU/µL SLO were compared. As observed earlier, the pristine membrane is highly insulating, with no observable redox peaks in the potential window studied. Incubation with SLO leads to a small increase in the anodic current (Supporting Information). This contrasts the pronounced anodic peak in the CV observed for preincubated vesicles for the same toxin concentration (Figure 5) where at +550 mV, the anodic current increased substantially from 0.7 to 2.5 µA. This also contrasts the results on the micropatterned bilayer patches where SLO incubation with existing membrane leads to a large redox signal.37 Here on the existing membrane at the same potential the current increases only from 0.6 to 0.9 µA, which is considerably small in both relative and absolute terms. In relative terms, incubation of an existing membrane increases the signal only by 50% whereas incubation of the vesicles yields a more than 300% increase. This shows that the supported PC

membrane on HT is likely to be far less mobile than the vesicles. A reduction in mobility is consistent with earlier observations that certain support layers such as alkanethiols reduce the lateral lipid diffusion and hence compromise the functions of membrane spanning inserts.41 For the in situ incubation approach, the small change in the CV makes it difficult to exactly determine the membrane permeability change. We hence used electrochemical impedance spectroscopy to further characterize the membrane properties based on its complex resistance. The obtained impedance spectra, separated into magnitude and phase difference for a pristine and incubated membrane, are shown in Figure 7. For both membranes the impedance magnitude (Z) decreased continuously with increasing frequencies, with Z digressing strongly at the higher end of the spectrum. The dispersion of the induced phase change was more complex. Two ranges of maximum digression were observed, between 100 and 1000 Hz and again at the high end of the spectrum. For a more quantitative analysis the impedance spectra were fitted to a simple equivalent circuit (inset in Figure 7). The circuit consists of a solution resistance (Rs) in series with a charge transfer resistance (RCT) and a constant phase element (CPE) connected in parallel. The fitting result shows a change in the membrane resistance (RCT) from 323 to 307 kΩ (about ca. 5%). The decrease in the charge transfer resistance is accompanied by the expected increase in the Q value of the CPE42 from 6.9 × 10-8 to 2.2 × 10-7 F. We believe that in the given system the 5% difference in the charge transfer resistance can be directly attributed to a 5% change in the membrane permeability. (41) Janshoff, A.; Steinem, C. Anal. Bioanal. Chem. 2006, 385, 433–451. (42) Vanwesting, E. P. M.; Ferrari, G. M.; Dewit, J. H. W. Electrochim. Acta 1994, 39, 899–910.

Toxin-Surface Interactions on Membranes

3. The Effect of Toxin, BSA and Surfactant on Redox Behavior of the HT Modified Electrode. A series of control experiments were conducted to verify that the specificity of the observed response is due to permeability changes in the lipid membrane. First the uncovered HT layer was incubated with SLO toxin. Toxin concentrations of 5 and 10 HU/µL, corresponding to molar concentrations of 63 and 125 nM, were studied. The resulting CVs show that compared to the pristine HT electrode surface, the current magnitudes for both the anodic and cathodic peak decreased consistently with increasing toxin concentrations (Supporting Information). For 10 HU/µL, both peak currents were reduced by about 50%, whereas for 5 HU/µL the current was reduced by ca. 11%. It is interesting to note that the electrode kinetics did not change for 5 HU/µL, with the peak separation remaining around 70 mV. Incubation with 10 HU/µL slowed down the electrode more severely, with the peak separation increased to 107 mV. The redox currents after incubation with 10 HU/µL were low, much lower than those observed on the permeated lipid membranes. We speculate that the hydrophobic nature of the HT SAM facilitates SLO toxin adsorption nonspecifically and reduces the surface area for redox reaction. The observed decrease in current for increasing concentration of toxins on HT electrodes contrasts the current increase for the lipid membrane system, further confirming the specificity of the perforation signal and highlighting the unique role that supporting membranes play in the study. In order to investigate the influence of nonspecific protein interactions at high concentrations with the lipid membrane, experiments were conducted with BSA on existing membranes and membranes generated from fusion of vesicles that were coincubated with BSA. The BSA concentration was 1 mg/mL, which is about 110-fold higher than the total protein concentration in the control experiments. Incubation of an existing membrane with BSA further reduced the current, indicating that process “repairs” the possible voids, pinholes, and other imperfections through blocking. The membranes obtained from vesicles coincubated with BSA are much less insulating, where anodic peak current was only reduced to 44% of the value on the HT electrode. We speculate that the presence of BSA during the incubation or the membrane fusion process strongly affects the membrane formation on HT. The average hydrodynamic diameter of the suspended vesicles as determined by optical light scattering is ca. 90 nm,23 while that of the BSA is ca. 7 nm.43 The BSA is hence diffusing much faster and could preferentially adsorb on the surface, creating “surface defects” in the form of BSA islands, obstructing the membrane formation. The effect of high concentration of BSA on HT modified electrode response was also studied. The direct incubation with BSA eliminated the cathodic current peak and shifted the anodic peak to a much higher potential with peak current reduced by ca. 27%. Apart from the structural change during SLO perforation, vesicles can be lysed by surfactants. We studied the influence of the nonionic surfactant Triton X-100 on the vesicle fusion/ membrane properties. Fusion of the Triton-incubated vesicle solution led to reduction of anodic currents to about 45% of that on the HT electrode. Compared to the membrane fused from (43) Axelsson, I. J. Chromatogr. 1978, 152, 21–32.

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intact vesicles, the Triton-treated membrane is apparently much less insulating. Nevertheless, the anodic peak current was lower than that of the membrane resulting from coincubation with BSA, and also lower than that from vesicles incubated with 1 HU/µL SLO. As no vesicle particles could be determined for the vesicle solution incubated with the Triton by using an optical particle sizer, we hence conclude that vesicle fragments, together with Triton, assemble onto the HT surface with fairly high surface coverage that results in reduction of anodic current. The effect of Triton on an existing membrane was also assessed since it is of particular value to the regeneration of the supporting layer. We found that addition of Triton onto the existing membrane on HT removed a significant amount of lipids from the surface but not completely. Compared to the fresh HT electrode, the peak current magnitude was reduced by about 35%, albeit with a much slower kinetics with anodic peak position shifting by 210 mV. It appears that the strong lysing activity of Triton X-100 on the suspended vesicles was relatively less effective for the preformed supported lipid membranes.

Conclusions The electrochemical behavior of three probes on two supporting layers has been evaluated for their suitability to study the permeability properties of supported lipid membranes manipulated by a pore-forming toxin. The support layer of hexyl thioctate and ferrocene carboxylic acid provided an ideal system for this purpose. The HT layer was highly permeable to the FCA probe while maintaining its fast kinetics. It further facilitated vesicle fusion on it to establish a stable insulating lipid membrane. Incubation of vesicles with SLO toxin and subsequent fusion on the HT modified electrode led to membranes that exhibit unique electrochemical characteristics. Over a concentration range of 0-10 HU/µL SLO, the electrochemical response showed a correlation to toxin concentration. High toxin concentrations increased the redox current magnitudes and enhanced the probe kinetics due to the increased number of pores in the membrane. Treating the existing PC membranes on the HT modified electrode with SLO resulted in a much reduced permeability change, likely due to the diminished fluidity for the hybrid structure. Exposure of the HT modified electrode to high concentrations of BSA induced nonspecific protein adsorption, diminishing the redox response and slowing down the kinetics of the probe. High concentration SLO toxin was also found to adsorb nonspecifically on HT and decrease the current. This behavior contrasts the current increase observed on the lipid membrane system obtained through fusion of SLO incubated vesicles. The HT/PC membrane system offers a unique platform to study and detect trace amounts of bacterial toxins via a specific perforation process. Future work will focus on further exploration of this mechanism for biosensing applications with supported lipid membranes. Supporting Information Available: Cyclic voltammograms for SLO, BSA, and Triton on the voltammetric response of FCA on the HT-modified Au electrodes. This information is available free of charge via the Internet at http://pubs.acs.org. LA704027C