Single europium-doped nanoparticles measure ... - Semantic Scholar

LETTERS PUBLISHED ONLINE: 9 AUGUST 2009 | DOI: 10.1038/NNANO.2009.200

Single europium-doped nanoparticles measure temporal pattern of reactive oxygen species production inside cells Didier Casanova1†, Ce´dric Bouzigues1†, Thanh-Lieˆm Nguyeˆn1†, Rivo O. Ramodiharilafy1, Latifa BouzhirSima1, Thierry Gacoin2, Jean-Pierre Boilot2, Pierre-Louis Tharaux3 and Antigoni Alexandrou1 * Low concentrations of reactive oxygen species, notably hydrogen peroxide (H2O2), mediate various signalling processes in the cell1,2. Production of these signals is highly regulated3 and a suitable probe is needed to measure these events. Here, we show that a probe based on a single nanoparticle can quantitatively measure transient H2O2 generation in living cells. The Y0.6Eu0.4VO4 nanoparticles undergo photoreduction under laser irradiation but re-oxidize in the presence of oxidants, leading to a recovery in luminescence. Our probe can be regenerated and reliably detects intracellular H2O2 with a 30-s temporal resolution and a dynamic range of 1–45 mM. The differences in the timing of intracellular H2O2 production triggered by different signals were also measured using these nanoparticles. Although the probe is not selective towards H2O2 , in many signalling processes H2O2 is, however, the dominant oxidant3–6. In conjunction with appropriate controls, this probe is a powerful tool for unravelling pathways that involve reactive oxygen species. Reactive oxygen species (ROS), and in particular H2O2 , have long been known for their microbial-killing role2. Moreover, it has been documented recently that different cell types produce H2O2 in much lower concentrations for signalling purposes1–3. The enzyme NADPH oxidase (NOx) produces the superoxide anion O22, which is then rapidly transformed into H2O2 (ref. 7). It is activated after stimulation by peptidic signals such as growth factors (platelet-derived4 and epidermal growth factor5 (PGDF and EGF)), cytokines8 and vasoconstricting peptides such as endothelin (ET-1)9–11. H2O2 regulates cell function by inhibiting phosphatases and activating kinases, transcription factors or Ca2þ signalling to control gene expression, cell growth or apoptosis1,2. It is anticipated that the cell response may be finely tuned by controlling the timing, amplitude and compartmentalization of H2O2 production3. Unravelling these pathways has, however, been hampered by the absence of an adequate sensor providing quantitative temporal and spatial subcellular information. The most widely used ROS and H2O2 probes, such as dichlorodihydrofluorescein diacetate, dihydrorhodamine and Amplex Red (Invitrogen), as well as other recently proposed alternatives12–16, rely on the formation of a fluorescent product and are therefore not reversible. They therefore detect the total amount of H2O2 generated but not the instantaneous concentration of H2O2. This effectively prohibits temporally resolved H2O2 measurements17. Although two reversible probes have been demonstrated18,19, they do not allow quantitative, intracellular detection at concentrations relevant for signalling (see Supplementary Discussion).

Silica-coated 20–40 nm Y0.6Eu0.4VO4 nanoparticles were spincoated and imaged individually using resonant Eu3þ excitation at 466 nm and detection at 617 nm (refs 20,21). Eu3þ ions show narrow absorption and emission lines corresponding to 4f 6–4f 6 dipole-forbidden intraconfiguration transitions largely independent of the host material (see Supplementary Fig. S1). On illumination, the luminescence of single nanoparticles decreases (Fig. 1a). Eu3þ ions are easily reduced (redox potential: Eu3þ/Eu2þ ¼ 20.35 V). In contrast to Eu3þ, Eu2þ ions show broadband absorption and emission due to dipole-allowed interconfiguration transitions 4f 65d1–4f 7 that depend on the host material, but Eu2þ absorption (emission) typically peaks around 330–430 nm (370–490 nm)22. The nanoparticle excitation and emission spectra in the range 270–590 nm show no absorption and emission lines before photobleaching, as expected, whereas after photobleaching the characteristic broad Eu2þ absorption and emission spectra appear (Fig. 1b). The decrease in luminescence is thus related to the photoreduction of Eu3þ to Eu2þ. The Eu2þ ions can be re-oxidized back to their initial Eu3þ state by various oxidants produced by cells (Fig. 1c; the normalized luminescence recovery DS/S0 is defined as the luminescence increase after normalization to 1 of the steady-state signal after photoreduction). However, the control experiments discussed below as well as data from the literature3–6 show that the dominant oxidant in many cell signalling processes is H2O2 (see Supplementary Discussion). O22 induces only a weak luminescence recovery due to its limited oxidation power23 and short lifetime7, which is even shorter inside cells in the presence of superoxide dismutases7. Intracellular NO concentrations are in the nM range24 and 2OCl is only produced in a reaction catalysed by myeloperoxidases in specialized cells such as neutrophils2. When adding varying concentrations C of H2O2 to the photoreduced nanoparticles, the luminescence recovery characteristics depend on C (Fig. 1d). In the absence of H2O2 (Fig. 1d, trace 0 mM), no deviation from the zero baseline is observed, indicating absence of spontaneous re-oxidation. Averages for collections of single nanoparticles are shown in Fig. 1c,d (see also Supplementary Fig. S2, which shows individual nanoparticle responses for C ¼ 17.8 mM). The excitation intensity and thus the relative importance of photoreduction versus oxidation determines the detectable oxidant concentration range. Low intensities allow detection of H2O2 concentrations down to 1 mM after averaging over a few nanoparticles (see Supplementary Fig. S3) and down to 2 mM for single particle detection, whereas higher intensities allow detection up to a few mM H2O2 (see Supplementary

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Laboratoire d’Optique et Biosciences, Ecole Polytechnique, CNRS, INSERM U696, 91128 Palaiseau, France, 2 Laboratoire de Physique de la Matie`re Condenseé, Ecole Polytechnique, CNRS, 91128 Palaiseau, France, 3 Paris-Cardiovascular Research Centre, INSERM U970, 56 rue Leblanc, 75015 Paris, France; † These authors contributed equally to this work. * e-mail: [email protected] NATURE NANOTECHNOLOGY | VOL 4 | SEPTEMBER 2009 | www.nature.com/naturenanotechnology © 2009 Macmillan Publishers Limited. All rights reserved.

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Figure 1 | Luminescence properties of silica-coated Y0.6Eu0.4VO4 nanoparticles spin-coated on silica coverslips. a, Luminescence decrease for a single nanoparticle as a function of illumination time (normalized to the value at t ¼ 0). b, Emission (blue and green curves; excitation at 310 nm) and excitation spectra (black and red curves; detection at 425 nm) of nanoparticles before (black and blue curves) and after (red and green curves) photobleaching (PB) at 3–5 kW cm22 for 30 s. c, Luminescence recovery after addition of various biological oxidants at t ¼ 0. [H2O2] ¼ 8.9 mM, [2OCl] ¼ 10+2 mM, [NO] ¼ 10 mM, cumulative [O22] 250 mM. d, Luminescence recovery after addition of various concentrations of H2O2 at t ¼ 0. Solid lines are fits using exponentials (equation (1)) except for the straight line at DS/S0 ¼ 0 for 0 mM H2O2. c and d show averages for a collection of single nanoparticles, N ¼ 20–30. e, Photoreduction/oxidation cycles for a single nanoparticle. Red arrows indicate addition of H2O2 and blue arrows indicate rinsing at the end of each cycle. [H2O2] ¼ 5 mM for the second and fourth cycle, [H2O2] ¼ 10 mM for all other cycles. Excitation intensity, 5 kW cm22; integration time, 0.5 s. Excitation at 466 nm, detection centred at 617 nm for all panels except b. Single nanoparticle measurements (a,c,d) were taken with an inverted microscope and ensemble measurements (b) with a spectrofluorometer.

Fig. S4). The photoreduction/re-oxidation cycles in Fig. 1e show that both the photoreduction and re-oxidation processes are reversible. This suggests that our nanoprobes can detect both rising and falling H2O2 concentrations. H2O2 detection is quantitative. The final luminescence value reached after recovery depends linearly on the H2O2 concentration in the 1–45 mM range (Fig. 2a). The recovery curves following a step-like addition of H2O2 can be fitted by exponentials with two concentration-dependent parameters, the characteristic time t (C) and the final value A(C ) (Fig. 2a,b): DS=S0 ¼ AðCÞ ð1 et=tðCÞ Þ

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concentration as a sequence of steps, and given that the monotonous relations t (C ) and A(C ) provide a unique solution for C(t) (Fig. 2d), equation (2) can be used to extract C(t), the instantaneous H2O2 concentration. The temporal resolution is determined by the time required for the measurement of (DS/S0)(t) and its first derivative and is typically 10–30 s. We then used this sensor to detect the H2O2 produced in signalling processes in living cells. In vascular cells, ET-1 and PDGF regulate contraction and migration by producing H2O2 (refs 4,9,10). We internalized the nanoparticles into mouse vascular smooth muscle cells (VSMCs) using pinocytosis and then photoreduced them. We verified the absence of nanoparticle toxicity to the cells using 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) tests (see Supplementary Fig. S5). The images in Figure 3a and b show, respectively, a white-light image of a VSMC and a fluorescence image of nanoparticles internalized in the same cell (see Supplementary Fig. S6 for a broader field-of-view image). In the absence of stimulation, the luminescence remains constant (see Supplementary Fig. S7). The addition of 1 mM exogenous H2O2 produced a luminescence recovery similar to that observed in vitro for 10 mM H2O2 due to intracellular H2O2 decomposition by catalases or other enzymes (see Supplementary Fig. S8), in agreement with ref. 19. The cells were then stimulated by ET-1 or PDGF at

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concentrations just above those producing a saturating response25,26. Activation of either the ET-1 or the PDGF receptor by their respective ligands leads to activation of the NOx enzyme9,11. Upon ET-1 stimulation, VSMCs almost instantaneously produce an intracellular oxidant, leading to luminescence recovery of the internalized nanoparticles, whereas no recovery is observed for a nanoparticle located outside a cell (Fig. 3c). Comparable behaviours of nanoparticles loaded at different cytosol locations reveal a homogeneous ROS distribution. Using the calibration of Fig. 2a and assuming that H2O2 is the dominant oxidant generated, we found an H2O2 production of C ¼ 13 mM in typically 25 min. The absence of recovery in the presence of apocynin, which degrades H2O2 (ref. 27) (Fig. 3d), or in the presence of catalase, which catalyses H2O2 decomposition to water (see Supplementary Fig. S9), confirms that the oxidative response in the cytosol is indeed exclusively due to H2O2. Similarly to ET-1, PDGF-BB stimulation induces the production of H2O2 (Fig. 4a), leading to a saturating concentration of C ¼ 7.1 mM after 30 min. In this case, however, a latency time of 5–10 min is observed. As for ET-1 stimulation, no luminescence recovery was observed in the presence of apocynin (Fig. 4a). To further confirm that the signal is due to H2O2 , we performed experiments after incubation with L-Name, a NO-synthase inhibitor: no significant differences were observed (Fig. 4a). The fit of the luminescence signal with a saturating power law was used to convert it into H2O2 concentration as a function of time based on the three-dimensional plot of Fig. 2d (Fig. 4b). After suppression of PDGF stimulation by rinsing, the luminescence decreases,

revealing intracellular H2O2 degradation (Fig. 4c). In this case, the decay time is limited by the temporal response of our probe, which is 7 min when the combined effect of laser-induced reduction and chemical oxidation results in reducing conditions for the nanoparticles and for the low excitation intensity used here. A notable difference between PDGF and ET-1 stimulation is the timing of H2O2 production. This difference may partly explain how the ET-1 and PDGF pathways can lead to distinct cell responses through production of the same secondary messenger (H2O2) and could rely on the mode of EGFR transactivation, which in turn can further activate NOx (ref. 5). ET-1 stimulation rapidly transactivates EGFR through activation of an EGFR ligand25,28, whereas PDGF-induced EGFR activation is based on EGFR/PDGFR heterodimerization29 and is probably slower. This implies that the observed H2O2 generation is at least partly due to NOx activation via the EGFR channel and that the fast response is possibly dominated by it. The timing of ROS production is thus a key element in signal transduction. This could constitute a general framework to understand how cells can produce a variety of responses with a limited number of signalling molecules. In conclusion, we have demonstrated a novel oxidant sensor based on oxido-reduction of Eu3þ ions in Y0.6Eu0.4VO4 nanoparticles and used it to quantify dynamically intracellular H2O2 concentrations produced under stimulation by two important regulators of cellular responses in the vascular system, PDGF and ET-1. These results constitute the first quantitative, time-resolved monitoring of ROS production and highlight the capability of the cell to


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Figure 3 | Intracellular H2O2 produced by VSMCs after ET-1 stimulation. a, White-light image of a VSMC. The cell nucleus can be distinguished on the left. The cell contour was enhanced with unsharp masking. b, Fluorescence image of Y0.6Eu0.4VO4 nanoparticles internalized in the same cell (excitation at 466 nm, detection centred at 617 nm) at t ¼ 30 min after ET-1 stimulation. Arrows indicate the nanoparticles analysed in c and d. No nanoparticles are observed inside the nucleus; defocused nanoparticles are located at other planes. c,d, Stimulation by ET-1 (270 nM at t ¼ 0): luminescence recovery of the three nanoparticles highlighted in b (blue, red and green curves) and signal of a nanoparticle located outside a cell in another experiment (black curve) (c); average signal of the three nanoparticles in b (blue curve) and average signal of three nanoparticles in the presence of 100 mM apocynin (black curve; the same results were obtained for three other cells) (d).

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Figure 4 | Intracellular H2O2 produced after PDGF stimulation. a, Luminescence recovery after stimulating VSMC with PDGF-BB at t ¼ 0 (100 ng ml21 or 4 nM; average data for N ¼ 5 cells, 4–5 nanoparticles per cell; blue circles), in the presence of 100 mM apocynin (N ¼ 5; green crosses), in the presence of 100 mM L-Name (N ¼ 4; red squares). Blue and red dashed lines are fits with DS/S0 ¼ At n/(Tn þ t n ) for A ¼ 0.33+0.01, T ¼ 19.3+0.8 min, n ¼ 3.1+0.3, and A ¼ 0.34+0.01, T ¼ 18.5+0.8 min, n ¼ 3.7+0.5, respectively. b, H2O2 concentration obtained numerically from the fit (dashed blue line) in a using the three-dimensional conversion plot in Fig. 2d. c, Luminescence decrease after rinsing off the buffer containing PDGF-BB and replacing it with HBSS/HEPES buffer (N ¼ 3 cells).

temporally regulate its response. The single particle detection capability can be used to assess cellular responses in asymmetric stimulation conditions. We can furthermore target these nanoprobes to cell compartments by appropriate functionalization30 to record their specific ROS response. Combined with pharmacological inhibition techniques, these nanoprobes indicate new perspectives for the deciphering of complex signalling pathways in a variety of biological systems.

Methods Luminescence characterization, excitation/emission spectra. All data in Figs 1 and 2 (except Fig. 1b) were obtained using nanoparticles spin-coated on silica 584

coverslips at low enough concentrations for individual observation. Ensemble excitation and emission spectra (see Supplementary Fig. S1 and Fig. 1b) were recorded using a Hitachi F-4500 spectrofluorometer. The spectra for assynthesized nanoparticles were obtained using nanoparticles in solution (see Supplementary Fig. S1). To record the spectra corresponding to photoreduced nanoparticles (Fig. 1b), we spin-coated the nanoparticle solution at a high concentration (300 times higher than for single particle observations) on a silica coverslip to form a quasi-uniform layer. A square window was then delimited on the coverslip and the 1-mm2 area inside the window illuminated with the microscope setup at 466 nm, 3–5 kW cm22 for 30 s by successive illumination of adjacent subareas. Excitation and emission spectra in the range expected for Eu2þ absorption and emission were recorded for the coverslip before and after 466-nm irradiation using the spectrofluorometer (Fig. 1b).



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Imaging procedures and materials. We used the 7F0,1–5D2 Eu3þ transition at 466 nm to excite the nanoparticles and detected their emission at the 5D0–7F2 Eu3þ transition consisting of two narrow lines at 615 and 619 nm (see Supplementary Fig. S1). As in ref. 20, we directly excited the Eu3þ ions at 466 nm rather than the vanadate matrix in the UV, which may be harmful for cells. The extinction coefficient at 466 nm was 24,000 M21 cm21 for a 30-nm 40%-doped nanoparticle and was sufficient for single particle observation. All images were recorded using an inverted microscope (Zeiss Axiovert 100) under wide-field illumination with the 465.8-nm line of an Arþ laser using a 63 or a 100, NA ¼ 1.4 oil-immersion objective (Zeiss) and a liquid-nitrogen-cooled charge-coupled device (CCD) (Princeton Instruments LN/CCD-400-PB, 400 1,340 pixels, back-illuminated) or an electron-multiplying CCD (Roper Scientific QuantEM:512SC). Nanoparticle luminescence was selected using a dichroic mirror (530DCXR, Chroma) and an emission filter (617/8M, Chroma). Measurements on spin-coated nanoparticles were performed in phosphate buffer, pH 7.4 at 20 8C. For all experiments (in vitro and inside cells), unless otherwise stated, the nanoparticles were photoreduced until a steady-state situation was reached using an excitation intensity of 1.6 kW cm22 at 466 nm for 250 s (integration time per image, 0.8 s). Illumination for 300 s at 0.3 kW cm22 without stimulation was used to ensure the stability of the luminescence. Oxidant detection was then performed (0.3 kW cm22; integration time, 2.8 s). We verified that the fraction of H2O2 molecules consumed to oxidize the Eu2þ ions was negligible, even for the lowest H2O2 concentrations considered here. H2O2 and 2OCl were added in the form of 30% and 12% aqueous solutions, respectively. O22 was obtained by adding 750 mM hypoxanthine at t ¼ 0 to a solution containing 25 munits ml21 xanthine oxidase and 100 units ml21 catalase (25 munits ml21 xanthine oxidase catalyses hypoxanthine into O22 at a rate of 50 mM min21 at 25 8C and pH 7.5). Catalase was added to ensure rapid decomposition of the H2O2 into water. NO was added in the form of a solution containing NO gas (excitation intensity for the recovery process: 1.6 kW cm22). For the reduction/oxidation cycles of Fig. 1e, the illumination was interrupted at the end of each cycle and the coverslip rinsed several times with phosphate buffer. The first bleaching cycle was not long enough (95 s) to reach the photobleached steady state (signal of 0.25), which was reached only after the second bleaching cycle. Therefore, the recovery response was also weaker (signal of 0.75 after the first recovery, compared with 0.9 after the third recovery). Cell imaging experiments were performed in HBSS/HEPES pH 7.4 buffer. All PDGF-BB stimulation experiments were performed at 20 8C. The final DS/S0 value of 0.33 obtained after PDGF-BB stimulation corresponds to a H2O2 concentration of 7.1 mM (Fig. 4a). The standard deviation for different cells (for different nanoparticles in the same cell) was 4 mM (3.6 mM), which is not much higher than the standard deviation of 2.2 mM for single nanoparticle responses in vitro. ET-1 stimulation experiments were performed either at 20 or 30 8C. The results in Fig. 3 were obtained at 30 8C. Similar results were obtained at 20 8C. N ¼ 11 cells in total. For control experiments with apocynin (L-Name), the cells were incubated with 100-mM concentrations for 30 (45) min at 37 8C in a humidified atmosphere containing 5% CO2. The same concentration (100 mM) was added in the observation buffer. Cell morphology was routinely controlled using white-light transmission before and after the experiment. Cells showing morphological changes after the experiment were not included in the statistics. We also used a trypan blue test to investigate cell viability after the full experimental process: pinocytic uptake of the nanoparticles, illumination for 250 s at 1.6 kW cm22 and stimulation by ET-1 for 30 min (N ¼ 9 cells). Eight out of nine cells were not stained by the dye (viable cells).

Received 24 March 2009; accepted 1 July 2009; published online 9 August 2009

References 1. Rhee, S. G. H2O2 , a necessary evil for cell signaling. Science 312, 1882–1883 (2006). 2. Bedard, K. & Krause, K.-H. The NOX family of ROS-generating NADPH oxidases: physiology and pathophysiology. Physiol. Rev. 87, 245–313 (2007). 3. D’Autreáux, B. & Toledano, M. B. ROS as signaling molecules: mechanisms that generate specificity in ROS homeostasis. Nature Rev. Mol. Cell Biol. 8, 813–824 (2007). 4. Sundaresan, M., Yu, Z.-X., Ferrans, V. J., Irani, K. & Finkel, T. Requirement for generation of H2O2 for platelet-derived growth factor signal transduction. Science 270, 296–299 (1995). 5. Bae, Y. S. et al. Epidermal growth factor (EGF)-induced generation of hydrogen peroxide. J. Biol. Chem. 272, 217–221 (1997). 6. Halliwell, B. & Cutteridge, J. M. C. Free Radicals in Biology and Medicine (Oxford Univ. Press, 1999). 7. Plonka, A. et al. Superoxide radical dismutation by copper proteins. J. Radioanal. Nucl. Chem. 101, 221–225 (1986).

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8. Lambeth, J. D. NOx enzymes and the biology of reactive oxygen. Nature Rev. Immunol. 4, 181–189 (2004). 9. Clempus, R. E. & Griendling, K. K. Reactive oxygen species signaling in vascular smooth muscle cells. Cardiovasc. Res. 71, 216–225 (2006). 10. Paravicini, T. M. & Touyz, R. M. Redox signaling in hypertension. Cardiovasc. Res. 71, 247–258 (2006). 11. Chen, C. H. et al. Reactive oxygen species generation is involved in epidermal growth factor receptor transactivation through the transient oxidization of Src homology 2-containing tyrosine phosphatase in endothelin-1 signaling pathway in rat cardiac fibroblasts. Mol. Pharmacol. 69, 1347–1355 (2006). 12. Soh, N. Recent advances in fluorescent probes for the detection of reactive oxygen species. Anal. Bioanal. Chem. 386, 532–543 (2006). 13. Hempel, S. L., Buettner, G. R., O’Malley, Y. Q., Wessels, D. A. & Flaherty, D. M. Dihydrofluorescein diacetate is superior for detecting intracellular oxidants: comparison with 20 ,70 -dichlorodihydrofluorescein diacetate, 5(and 6)-carboxy20 ,70 -dichlorodihydrofluorescein diacetate, and dihydrorhodamine 123. Free Radic. Biol. Med. 27, 146–159 (1999). 14. Miller, E. W., Albers, A. E., Pralle, A., Isacoff, E. Y. & Chang, C. J. Boronate-based fluorescent probes for imaging cellular hydrogen peroxide. J. Am. Chem. Soc. 127, 16652–16659 (2005). 15. Miller, E. W., Tulyathan, O., Isacoff, E. Y. & Chang, C. J. Molecular imaging of hydrogen peroxide produced for cell signaling. Nature Chem. Biol., 3 263–267 (2007). 16. Lee, D. et al. In vivo imaging of hydrogen peroxide with chemiluminescent nanoparticles. Nature Mater. 6, 765–769 (2007). 17. Seshiah, P. N. et al. Angiotensin II stimulation of NAD(P)H oxidase activity: upstream mediators. Circ. Res. 91, 406–413 (2002). 18. Wolfbeis, O. S., Duerkop, A., Wu, M. & Lin, Z. Europium ion-based luminescent sensing probe for hydrogen peroxide. Angew. Chem. Int. Ed. 41, 4495–4498 (2002). 19. Belousov, V. V. et al. Genetically encoded fluorescent indicator for intracellular hydrogen peroxide. Nature Meth. 3, 281–286 (2006). 20. Beaurepaire, E. et al. Functionalized fluorescent oxide nanoparticles: artificial toxins for sodium channel targeting and imaging at the single molecule level. Nano Lett. 4, 2079–2083 (2004). 21. Casanova, D., Giaume, D., Gacoin, T., Boilot, J.-P. & Alexandrou, A. Optical in situ size determination of single lanthanide-ion doped oxide nanoparticles. Appl. Phys. Lett. 89, 253103 (2006). 22. Dorenbos, P. Anomalous luminescence of Eu2þ and Yb2þ in inorganic compounds. J. Phys. Condens. Matt. 15, 2645–2665 (2003). 23. Sawyer, D. T. & Valentine, J. S. How super is superoxide? Acc. Chem. Res. 14, 393–400 (1981). 24. Hall, C. N. & Attwell, D. Assessing the physiological concentration and targets of nitric oxide in brain tissue. J. Physiol. 586, 3597–3615 (2008). 25. Chansel, D. et al. Heparin binding EGF is necessary for vasospastic response to endothelin. FASEB J. 20, E1368–E1381 (2006). 26. Rosenkranz, S. et al. Inhibition of the PDGF receptor by red wine flavonoids provides a molecular explanation for the ‘French paradox’. FASEB J. 16, 1958–1960 (2002). 27. Heumu¨ller, S. et al. Apocynin is not an inhibitor of vascular NADPH oxidases but an antioxidant. Hypertension 51, 211–217 (2008). 28. Daub, H., Weiss, F. U., Wallasch, C. & Ullrich, A. Role of transactivation of the EGF receptor in signalling by G-protein-coupled receptors. Nature 379, 557–560 (1996). 29. Saito, Y., Haendeler, J., Hojo, Y., Yamamoto, K. & Berk, B. C. Receptor heterodimerization: essential mechanism for platelet-derived growth factorinduced epidermal growth factor receptor transactivation. Mol. Cell. Biol. 21, 6387–6394 (2001). 30. Casanova, D. et al. Counting the number of proteins coupled to single nanoparticles. J. Am. Chem. Soc. 129, 12592–12593 (2007).

Acknowledgements We thank G. Mialon and M. Moreau for nanoparticle synthesis.

Author contributions D.C., C.B. and T.-L.N. contributed equally. D.C., C.B., P.-L.T. and A.A. conceived and designed the experiments. D.C., C.B. and T.-L.N. performed the experiments. D.C., C.B., T.-L.N. and A.A. analysed the data. R.O.R. and L.B.-S. contributed the cell cultures and data on nanoparticle toxicity. T.G. and J.-P.B. contributed the nanoparticles. All authors discussed the results. C.B. and A.A. co-wrote the paper.

Additional information Supplementary information accompanies this paper at www.nature.com/ naturenanotechnology. Reprints and permission information is available online at http://npg.nature.com/reprintsandpermissions/. Correspondence and requests for materials should be addressed to A.A.


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