Anal. Chem. 2008, 80, 9635–9641
Triangulation Mapping of Oxidative Bursts Released by Single Fibroblasts by Amperometry at Microelectrodes Christian Amatore,* Ste ´ phane Arbault, and Marie Erard† Laboratoire Pasteur, Ecole Normale Supe´rieure, CNRS, UPMC Universite´ Paris 06, De´partement de Chimie, 24 Rue Lhomond, 75005, Paris, France It has been previously established that a lesion created by a microcapillary in the membrane of a single aerobic cell (from skin or immune origin) was sufficient to induce a local membrane depolarization and the ensuing release of oxidative bursts. Their kinetic and quantitative features reveal the activity of cell constitutive enzymes, namely, NADPH oxidases and NO synthases, prone to produce rapidly reactive oxygen and reactive nitrogen species. Until now, the spatial resolution provided by microelectrodes has been exploited in this context to characterize the chemical composition of oxidative bursts at several cell types with high collection efficiency. In the present work, spatial features of the oxidative bursts from single human fibroblasts were investigated using a step-by-step geometrical mapping approach. The spatial locations of cell active zones and of the extent of the activated area, when a cell membrane was stressed by a microcapillary’s tip of 1-µm radius, have been addressed. On cells of large dimensions such as fibroblasts, ROS and RNS emission originated from a disk surface of the membrane limited to ∼15-µm radius around the ∼1-µm hole created by the microcapillary. This experimental result was rationalized through a simple physicochemical model designed to portray the extent of the membrane activated area due to ion concentration variations resulting from the pinhole channel created across the cell membrane. This is consistent with the fact that the activation of constitutive enzymatic complexes (NOX and NOS) is hypothesized to be a consequence of local variations of ion concentrations such as K+, Na+ or possibly Ca2+. Our results showed that the calculated area near the cell membrane where the ion concentration gradients are significant was equivalent to the area of species release measured experimentally. Aerobic cells actively produce several chemical reactive species, namely, reactive oxygen (ROS) and reactive nitrogen species (RNS), when their integrity is threatened by environmental hazards such as infectious entities (e.g. virus, bacteria), xenobiotics (e.g. water or air pollutants), or physical agents (e.g. high* To whom correspondence should be addressed. E-mail:
[email protected]. † Present address: Laboratoire de Chimie Physique, UMR 8000, Universite´ Paris Sud, F-91405 Orsay, France/CNRS F-91405 Orsay, France. 10.1021/ac801269e CCC: $40.75 2008 American Chemical Society Published on Web 11/08/2008
energy radiation, mechanical stress). The ensuing increase of oxidizing species concentrations in cells or biological fluids is generally compensated by endogenoussor by producedsreducing substances (e.g., vitamins, glutathione, catalase, superoxide dismutase). When the balance between ROS/RNS and antioxidants is not equilibrated, cells present a physiological situation called oxidative stress and are prone to consequently evolve toward abnormal phenotypes or death processes.1-5 We and other authors have shown that electrochemical analyses with microelectrodes constitute highly efficient methodologies to investigate qualitatively and quantitatively the production or the release of reactive oxygen and nitrogen species at the level of a single eukaryote cell.6-16 In particular, we have reported that cells from the dermis of skin, i.e., fibroblasts, release oxidative bursts when stimulated by a fast pricking of their membrane with a sealed microcapillary (e1-µm radius).6-8 This stimulus is supposed to induce a localized activation of enzymatic systems through local ionic changes and consequent membrane depolarization. For the detection, chemical characterization and quantification of ROS and RNS released by the activated area, a platinized carbon fiber microelectrode of ∼10-µm diameter is placed at micrometric distances above the cell by way of micromanipulators. This configuration affords an analysis in real time of the chemical blend emitted as a burst by the cell. We thus (1) Halliwell, B.; Gutteridge, J. M. C. Free Radicals in Biology and Medicine, 4th ed.; Oxford University Press: New York, 2007. (2) Finkel, T.; Holbrook, N. J. Nature 2000, 408, 239–247. (3) Finkel, T. Nat. Rev. Mol. Cell Biol. 2005, 6, 971–976. (4) Nathan, C.; Shiloh, M. U. Proc. Natl. Acad. Sci. U. S. A. 2000, 97, 8841– 8848. (5) Pacher, P.; Beckman, J. S.; Liaudet, L. Physiol. Rev. 2007, 87, 315–424. (6) Arbault, S.; Pantano, P.; Sojic, N.; Amatore, C.; Best-Belpomme, M.; Sarasin, A.; Vuillaume, M. Carcinogenesis 1997, 18, 569–574. (7) Arbault, S.; Pantano, P.; Jankowski, J. A.; Vuillaume, M.; Amatore, C. Anal. Chem. 1995, 67, 3382–3390. (8) Amatore, C.; Arbault, S.; Bruce, D.; De Oliveira, P.; Erard, M.; Vuillaume, M. Faraday Discuss. 2000, 319–333. (9) Amatore, C.; Arbault, S.; Bouton, C.; Coffi, K.; Drapier, J. C.; Ghandour, H.; Tong, Y. H. ChemBioChem 2006, 7, 653–661. (10) Privat, C.; Stepien, O.; David-Dufilho, M.; Brunet, A.; Bedioui, F.; Marche, P.; Devynck, J. Free Radical Biol. Med. 1999, 27, 554–559. (11) Isogai, Y.; Tsuyama, T.; Osada, H.; Iizuka, T.; Tanaka, K. FEBS Lett. 1996, 380, 263–266. (12) Malinski, T.; Taha, Z. Nature 1992, 358, 676–678. (13) Hill, H. A. O.; Tew, D. G.; Walton, N. J. FEBS Lett. 1985, 191, 257–263. (14) Xue, J.; Ying, X. Y.; Chen, J. S.; Xian, Y. H.; Jin, L. T.; Jin, J. Anal. Chem. 2000, 72, 5313–5321. (15) Kubant, R.; Malinski, C.; Burewicz, A.; Malinski, T. Electroanalysis 2006, 18, 410–416. (16) Isik, S.; Schuhmann, W. Angew. Chem., Int. Ed. 2006, 45, 7451–7454.
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showed previously that the oxidative response of fibroblasts in culture results from the initial production of nitric oxide radical (NO•) and superoxide anion (O2•-) by enzymatic systems, namely, NO synthases (NOS) and NADPH oxidases (NOX), of the cell membrane. According to analyses at several selected potentials, one may identify and quantify individually the release of hydrogen peroxide (H2O2), peroxynitrite (ONOO-), nitrogen monoxide (NO•), and nitrites (NO2-) by each fibroblast, all these species being the follow-up products of the fluxes of NO• and O2•- primarily produced by the cell.17,18 In a previous study,8 we investigated the collection efficiency of ROS and RNS release at single fibroblasts by platinized carbon microelectrodes. We have reported that the collection of the cell response was quantitative at 5-µm distance from the cell membrane and closer, thus allowing the quantitative characterization of the chemical blend emitted. However, two important issues regarding the localization and the spatial extent of the cell area leading to the emission of ROS and RNS have not been investigated yet. The first one concerns the surface sites of plasma membrane where the emission of the electroactive species takes place. In this work, we address the question of a localized emission of the ROS and RNS in contrast with the possibility of a global emission by the whole membrane of the cell. The second question is aimed to address the correlation of the spatial extent of the cell membrane area responsible for response with that (a few micrometers squared at most) directly injured and stimulated by the microcapillary. In order to answer these two crucial questions, we propose in the following a triangulation-based approach compatible with the time duration of oxidative bursts. This was achieved by monitoring the cell responses at different fixed locations of the electrode versus the cell membrane instead of a continuous and automatized mapping such like in SECM.16,19-21 The methodological approach presented below presents several advantages in our particular situation. Indeed, due to the large cellular surface of fibroblasts (30 µm × 80 µm, in average) compared to the microelectrode’s tip size (10-12 µm diameter), a continuous scanning is less essential than the ability to cover large distances, up to 80 µm. Besides, the time scale of each cell spiked response (20-s half-width in average) is only compatible with a single point measurement: in this situation, displacement of the microelectrode’s tip over the cell surface while keeping the ability to measure equivalent signals is very unlikely since the oxidative bursts have amplitudes that rapidly decay with time. Consequently, measurements at different selected locations, referenced to the zone of membrane where a stress is applied by a microcapillary, have been compared over a statistically significant set of cells. This offered a true characterization of zones activated by a micrometer-scale membrane puncture. (17) Amatore, C.; Arbault, S.; Bruce, D.; De Oliveira, P.; Erard, M.; Sojic, N.; Vuillaume, M. Analusis 2000, 28, 506–517. (18) Amatore, C.; Arbault, S.; Bruce, D.; De Oliveira, P.; Erard, M.; Vuillaume, M. Chem. Eur. J. 2001, 7, 4171–4179. (19) Schulte, A.; Schuhmann, W. Angew. Chem., Int. Ed. 2007, 46, 8760–8777. (20) Gao, N.; Wang, X. L.; Li, L.; Zhang, X. L.; Jin, W. R. Analyst 2007, 132, 1139–1146. (21) Diakowski, P. M.; Ding, Z. F. Phys. Chem. Chem. Phys. 2007, 9, 5966– 5974.
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EXPERIMENTAL SECTION Chemicals. All chemical reagents were purchased from Sigma-Aldrich (St. Louis, MO). Phosphate buffer, PBS, was used as buffer (137 mM NaCl, 2,7 mM KCl, 10 mM H2PO4; pH 7.4) throughout all experiments and was prepared by dissolving tablets in ultrapure water (Milli-Q system; Millipore). Cell Culture. The fibroblasts used in this study were obtained from a normal human cell strain established from a skin biopsy. They have been supplied to us by the laboratory of Dr. A. Sarasin (CNRS UPR 2169, Institut Gustave Roussy, Villejuif, France). They were grown in MEM F12 medium (Gibco BRL) supplemented with fetal calf serum (10%) and antibiotics (penicillin 10 000 units/ mL, streptomycin 10 mg/mL, fungizone 0.25 mg/mL) in an incubator (5% CO2, 37 °C). Monolayer of confluent cells were harvested by trypsination (trypsin-EDTA). The 1000-2000 cells were then resuspended in each plastic Petri dish (3.5-cm diameter; Costar 3035) and analyzed 24-48 h later; meanwhile, cells spontaneously adhered to the Petri dish surface. Just before the experiments, cells were washed three times with PBS, and the measurements were performed with isolated cells to avoid any interference between products possibly released by the neighboring cells during the oxidative bursts. Platinized Carbon Fiber Microelectrodes. Carbon fiber microelectrodes (10-µm diameter, Thornel Carbon Fibers, Cytec Engineered Materials, Greenville, SC) were constructed as described previously7 and backfilled with mercury for electrical contact. Electrode tips were polished at a 45° angle on a diamond particle whetstone microgrinder (model EG-4, Narishige, London, UK) for 2 min before platinization. The highly electrocatalytic porous black platinum coating was grown on the polished carbon fiber surface by reducing a solution of hydrogen hexachloroplatinate(IV) in PBS in the presence of lead(II) acetate trihydrate at -60 mV.7,22,23 The process was stopped when the electrical charge delivered reached 80-90 µC. The average diameter of the apparent area of the microelectrode tip was 12 µm. The microelectrodes were stored in distilled water at 4 °C for several days until use. Only electrodes giving a very stable amperometric baseline current after polarization at 450 mV versus saturated sodium chloride calomel electrode (SSCE) under in vitro conditions were used for cell measurements. Amperometric Measurements. The experiments were performed on the stage of an inverted microscope (Axiovert 135, Zeiss), where the Petri dish was positioned with cells to study. The movements of the platinized carbon fiber microelectrode and of the sealed glass microcapillary (1-mm glass rods, GR100-10, Clark Instruments, GB; pulled with a PB7 puller, Narishige, Japan; tip diameter below 1 µm) were controlled with two micromanipulators (MHW-103, Narishige, Japan) with a precision of 0.5 µm. The tip of the microcapillary (radius below 1 µm) was set under optical control on the plasma membrane at the place where the stimulus was to be applied. According to the micropositioner precision (
