An aryleneethynylene fluorophore for cell membrane staining

Biochimica et Biophysica Acta 1818 (2012) 2808–2817

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An aryleneethynylene fluorophore for cell membrane staining Antonio Cardone a, Francesco Lopez b, Francesco Affortunato a, Giovanni Busco c, Aldebaran M. Hofer d, Rosanna Mallamaci c, Carmela Martinelli a, Matilde Colella c,⁎⁎, Gianluca M. Farinola e,⁎ a

Consiglio Nazionale delle Ricerche, Istituto di Chimica dei Composti OrganoMetallici, UOS di Bari, Via Orabona, 4-70125 Bari, Italy Dipartimento di Agricoltura, Ambiente ed Alimenti (DIAAA) and CSGI, Università degli Studi del Molise, I-86100 Campobasso, Italy Dipartimento di Bioscienze, Biotecnologie e Scienze Farmacologiche, Università degli Studi di Bari “Aldo Moro”, Via Orabona, 4-70125 Bari, Italy d VA Boston Healthcare System and the Department of Surgery, Brigham and Women's Hospital and Harvard Medical School, 1400 VFW Parkway, West Roxbury, MA 02132, USA e Dipartimento di Chimica, Università degli Studi di Bari “Aldo Moro”, Via Orabona, 4-70125 Bari Italy b c

a r t i c l e

i n f o

Article history: Received 27 December 2011 Received in revised form 2 June 2012 Accepted 16 June 2012 Available online 27 June 2012 Keywords: Aryleneethynylene Fluorescent marker pH-insensitive Membranes Cross-coupling

a b s t r a c t The use of an amphiphilic aryleneethynylene fluorophore as a plasma membrane marker in fixed and living mammalian cells and liposome model systems is demonstrated. We show here that the optical properties of the novel dye are almost independent on pH, in the range 5.0–8.0. Spectroscopic characterization performed on unilamellar liposomes ascertained that the fluorescence intensity of the aryleneethynylene fluorophore greatly increases after incorporation in lipidic membranes. Experiments performed on different mammalian cells demonstrated that the novel membrane marker exhibits fast staining and a good photostability that make it a suitable tool for live cell imaging. Importantly, the aryleneethynylene fluorophore was also shown to be a fast and reliable blue membrane marker in classical multicolor immunofluorescence experiments. This study adds new important findings to the recent exploitation of the wide class of aryleneethynylene molecules as luminescent markers for biological investigations. © 2012 Elsevier B.V. All rights reserved.

1. Introduction In the last few years, great research efforts have been devoted to the development of novel fluorescent probes to selectively stain proteins and monitor organelle dynamics and ion homeostasis in living cells and organisms. In particular, since many cell functions are regulated through dynamic modifications of plasma membrane, different fluorescent dyes have been developed to sense changes in the membrane environment [1], polarization and spreading of cells [2], changes in membrane fluidity [3] and lipid rafts [4], just to name a few. In the choice of fluorescent markers, molar absorptivity, quantum yield of fluorescence, chemical stability, low toxicity and low cost are crucial criteria. Among the most serious drawbacks of many commonly used fluorescent probes there are susceptibility to photobleaching [5] and pH sensitivity [6].

Abbreviations: INS1-E cells, insulin secreting beta cells; BHK-21 cells, Baby Hamster Kidney cells; DMSO, dimethyl sulfoxide; DDAB, dimethyl–dioctadecyl ammonium bromide; HEPES, (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid); DPH, 1,6-Diphenyl-1,3,5hexatriene; DHPE Oregon Green, 1,2-Dihexadecanoyl-sn-Glycero-3-Phosphoethanolamine Oregon Green ⁎ Corresponding author. Fax: +39 080 5442129. ⁎⁎ Corresponding author. Fax: +39 080 5443388. E-mail addresses: [email protected] (M. Colella), [email protected] (G.M. Farinola). 0005-2736/$ – see front matter © 2012 Elsevier B.V. All rights reserved. doi:10.1016/j.bbamem.2012.06.011

In order to minimize pH sensitivity, a number of structural modifications have been used. In the case of fluorescein-related compounds, for example, a highly fluorescent monoanionic fluorophore [7] has been obtained by replacing the carboxylate with the methyl group or, alternatively, by functionalizing the molecules with fluorine atoms [8]. These structural changes resulted in improved fluorescence efficiency at low pH and significantly increased the photostability of the fluorescein derivatives. To date, a wide variety of light emitting compounds has been investigated as fluorescent probes tailoring molecular structures and emission wavelengths for specific labeling in different biological applications. Due to their efficient fluorescence and good stability [9], aryleneethynylene oligomers and polymers, upon proper functionalization, have been widely investigated for sensing applications of biological interest [10,11]. Examples include carbohydrate-functionalized poly(phenyleneethynylene)s (PPEs) for detection of Escherichia coli by multivalent interactions [12], folate-functionalized PPE as fluorescent contrast agents to detect cancer cells in vitro [13], cationic PAE polyelectrolytes for monitoring phospholipase A2 activity [14], sulfonated and biotinylated PPE-coated streptavidin-derivatized polystyrene microspheres as platform for the detection of DNA hybridization [15], pendant amine-functionalized PPE for detection of Cy-5-labeled oligonucleotides [16], biotin-functionalized PPE and streptavidin-derivatized microspheres as model systems for the recognition that occurs in cells and bacteria [17], PPEs-based fluorogenic probe for proteases [18]. Despite their efficient and broadly tuneable emission properties, chemical robustness and easy functionalization, to the best of our

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knowledge aryleneethynylene oligomers have never been investigated as fluorescent markers for biological applications, and only recently polyarylenethynylenes have been proven to be suitable as fluorescent tools for live and fixed cell imaging [19,20]. The restriction of the use of aryleneethynylenes as biological markers has been most probably determined by the high energy UV excitation necessary to excite these compounds. It is well known that high energy UV radiation has several general disadvantages in live cell imaging [21], such as increased phototoxicity, weak penetration into multicellular preparations due to high absorption of the short wavelength light, high scattering and contamination from tissue autofluorescence. As far as the phototoxicity is concerned, the group of Whitten has extensively worked on poly(phenylene ethyneylene) (PPE)-based cationic conjugated polyelectrolytes (CPEs) and cationic phenyleneethynylene oligomers (OPEs) [22–24]. These authors showed that most of these compounds (bearing cationic groups) exhibit dark and light-activated biocidal activity. As described for other organic dyes, the light-activated biocidal action of these compounds has been attributed to the generation of singlet oxygen upon exposure to UV–visible light. This potential weakness of aryleneethynylenes has turned in a strong point in favour of their use in live cell imaging experiments when novel microscopic techniques, such as two-photon excitation microscopy, have become available to researchers [19,20]. In fact, by enabling the excitation of fluorophores by the combination of the energy of two near-simultaneous double-wavelength photons, two-photon microscopy eliminates some of the problems associated with excitation spectrum in UV range. In connection with our recent research on functionalized organic semiconductors for advanced sensing applications [25,26] also covering phenyleneethynylene structures [11,27–29], we report herein a study on the use of an ad hoc synthesized amphipilic aryleneethynylene fluorophore suitable for liposome and cell membrane staining, further confirming that these luminescent molecules, if opportunely functionalized, can be conveniently used as a novel class of membrane markers for a wide range of biological applications. 2. Materials and methods 2.1. Materials and solutions All chemicals were purchased from SIGMA Aldrich, Alfa Aesar or Acros and used without further purification. L-α-phosphatidylcholine (Egg, Chicken) 99% was from Avanti Polar Lipids (Alabaster, AL). DHPE Oregon Green was purchased from Life technologies. THF was distilled from sodium and benzophenone under a nitrogen atmosphere. DMF and Et3N were distilled from 4 Å molecular sieves under a nitrogen atmosphere. All solvents were freshly distilled immediately prior to use. Biological experiments were performed with a Ringer's solution containing (in mM): 121 NaCl, 2.4, K2HPO4, .4 KH2PO4, 1.2 CaCl2, 1.2 MgCl2, 5.5 glucose, 10 HEPES (pH 7.4). When DMSO or EtOH were used, the final solvent concentration in Ringer's solution never exceeded 0.01% or 0.1%, respectively. 2.2. Methods Column chromatography was performed using silica gel 60, 40–63 μm from Merck. Merck Silica gel 60F254 aluminium sheets were used for analytical TLC. FT-IR spectra were measured on a Perkin-Elmer 1710 spectrophotometer using dry KBr pellets. 1H and 13C NMR spectra were recorded in CDCl3 or DMSO-d6 on a Bruker AM 500 spectrometer at 500 MHz and 125 MHz, respectively. The residual CHCl3 or DMSO signal at δ=7.24 (or 2.49) ppm and the CDCl3 (or DMSO-d6) signal at δ=77.0 (or 39.5) ppm were used as the standards for 1H NMR and 13C NMR spectra, respectively. MS spectra were recorded on a Shimadzu GCMS-QP 5000 spectrometer. Elemental analyses were done by a Carlo Erba CHNS-O EA1108-Elemental Analyzer. Melting points were determined on a

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Gallenkamp capillary melting point apparatus. UV–vis spectra were recorded on a Shimadzu UV-2401 PC spectrophotometer. Excitation and emission spectra were recorded with a Varian Cary Eclipse Fluorimeter. 2.3. Synthesis of intermediate compounds 2.3.1. Trimethyl-(4-octadecyloxy-phenylethynyl)-silane (1) 1-Iodo-4-(octadecyloxy)benzene [30] (3.03 g, 6.42 mmol), PdCl2(PPh3)2 (0.09 g, 0.13 mmol), CuI (0.013 g, 0.07 mmol) and dry Et3N (60 mL) were mixed at room temperature in a 100 mL three necked round bottom flask under nitrogen atmosphere and stirred for 10 min. Then, trimethylethynyl-silane (0.98 mL, 7.06 mmol) was added and the resulting mixture stirred overnight at room temperature. After removal of the reaction solvent at reduced pressure, petroleum ether was added (100 mL) and the organic layer washed with aq. HCl 5% (50 mL) and dried over anhydrous Na2SO4. After evaporation of the solvent, the crude product was purified by crystallization from CH3CN, to yield 1 as a white solid powder (2.35 g, yield 83%). Mp 48–49 °C (CH3CN). 1H NMR (CDCl3) δ: 7.38 (d, J=8.8 Hz, 2H), 6.80 (d, J=8.8 Hz, 2H), 3.94 (t, J=6.6 Hz, 2H), 1.77 (quint, J=6.6 Hz, 2H), 1.44 (quint, J= 6.6 Hz, 2H), 1.26 (s, 28H), 0.88 (t, J=6.6 Hz, 3H), 0.24 (s, 9H). 13C NMR (CDCl3) δ: 159.32, 133.42, 114.92, 114.29, 105.29, 92.25, 68.02, 31.92, 29.69, 29.65, 29.58, 29.55, 29.37, 29.16, 25.99, 22.69, 14.13, 0.07. FT-IR (KBr): 2919 (31), 2849 (36), 1607 (42), 1245 (37), 868 (39), 840 (38) cm−1. MS (EI, 70 eV): m/z (%)=442 (M+, 10), 355 (10), 220 (13), 206 (33), 193.(51), 190 (54), 175 (100), 166 (11), 73 (11). Anal. Calcd for C29H50OSi: C, 78.66; H, 11.38. Found C, 78.53; H, 11.51%. 2.3.2. Sodium 3,3′-(2,7-diiodo-9H-fluorene-9,9-diyl)dipropane-1-sulfonate (2) A solution of 2,7-diiodofluorene (3 g, 7.18 mmol) and benzyltriethylammonium chloride (0.16 g, 0.71 mmol) in DMSO (80 mL) was bubbled with a nitrogen flow for 20 min in a 250 mL three necked round bottom flask; then aq. NaOH 50% (1.27 mL) was added and the mixture stirred for 30 min at room temperature. A solution of 1,3-propanesultone (2.19 g, 17.94 mmol) in DMSO (20 mL) was slowly added to the reaction mixture, and the reaction was stirred overnight at room temperature. After addition of acetone, the precipitate was filtered and washed with acetone. The crude solid was purified by crystallization with water/acetone to yield the pure product 2 as a white solid (3.04 g, yield 60%). Mp>350 °C (H2O/acetone). 1H NMR (DMSO-d6) δ: 7.82 (s, 2H), 7.70 (d, J=8.0, 2H), 7.66 (d, J=8.0, 2H), 2.17 (t, J=8.1, 4H), 1.98 (t, J=8.3, 4H), 0.71 (quint, J=4.9, 4H). 13C NMR (DMSO-d6) δ: 151.81, 139.34, 135.90, 131.59, 122.23, 93.97, 55.16, 51.62, 38.43, 19.63. FT-IR (KBr): 1641 (91), 1449 (86), 1409 (91), 1178 (61), 1051 (70), 816 (87) cm−1. Anal. Calcd for C19H18I2Na2O6S2: C, 32.31; H, 2.57; S, 9.08. Found C, 32.20; H, 2.61; S, 9.20%. 2.4. Synthesis of the oligomer O1 2.4.1. Sodium 3-[2,7-bis-(4-octadecyloxy-phenylethynyl)-9-(3sodiumsulfonate-propyl)-9H-fluoren-9-yl]-propane-1-sulfonate (O1) Dry DMF (25 mL), dry THF (10 mL), reagent 1 (0.92 g, 2.08 mmol), reagent 2 (0.7 g, 1.04 mmol), Pd(PPh3)4 (0.06 g, 0.05 mmol) and Ag2O (0.48 g, 2.08 mmol) were added in a 100 mL three necked round bottom flask under nitrogen atmosphere, and the resulting mixture was stirred overnight at 60 °C. After cooling to room temperature, the reaction mixture was filtered through celite to remove Ag2O, then by adding acetone a yellow-brown precipitate was collected. A recrystallization from MeOH/acetone yielded the pure product O1 as an orange solid (0.78 g, yield 63%). Mp 220–230 °C (MeOH/acetone). 1H NMR (DMSO-d6) δ: 7.88 (d, J = 7.75 Hz, 2H), 7.61 (s, 2H), 7.52 (tlike, J = 8.4 Hz, 6H), 6.96 (d, J = 8.7 Hz, 4H), 4.0 (t, J = 6.3 Hz, 4H), 2.30–1.90 (bs, 8H), 1.71 (quint, J = 6.9 Hz, 4H), 1.50–1.40 (bs, 60H), 0.95–0.60 (bs, 10H). 13C NMR (DMSO-d6) δ: 158.88, 150.46, 140.03, 132.98, 132.90, 125.59, 121.64, 120.56, 114.71, 114.16, 90.05, 88.78, 67.54,

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54.86, 51.73, 31.23, 28.95, 28.69, 28.63, 28.55, 25.41, 22.03, 13.88. FT-IR (KBr): 2921 (32), 2851 (38), 1602 (46), 1508 (41), 1246 (38), 1173 (40), 1051 (43), 829 (46), cm−1. Anal. Calcd for C71H100Na2O8S2: C, 71.56; H, 8.46; S, 5.38. Found: C, 71.72; H, 8.30; S, 5.29%. 2.5. Spectroscopic measurements on liposomes Large unilamellar liposomes were prepared by reverse phase evaporation, according to the method described by Szoka [31]. Phosphatidylcholine and dimethyl-dioctadecyl ammonium bromide (DDAB) were mixed (6.5:3.5 molar ratio) in 20 mM HEPES (pH 7.2) to achieve a final phospholipid concentration of 15 mg/mL (i.e. 20 mM) [32]. Liposomes were then prepared by stepwise extrusion through 0.1 μm and 0.05 μm polycarbonate membranes. This method typically yields homogeneous liposomes with a mean diameter of 80 nm as assessed by Dynamic Light Scattering (DLS). Steady-state fluorescence measurements were performed using a Varian Eclipse spectrofluorimeter in a 1-cm quartz fluorescence cuvette, at 25 °C after an incubation time of 10 min. The excitation and emission slit widths were 5 nm. 2.6. Fluorescence studies on mammalian cells 2.6.1. Cell culture BHK-21 and HeLa cells were purchased from ATCC, INS-1E were a generous gift from Prof. C. Wollheim [33], cardiac fibroblasts form neonatal rats were obtained as described elsewhere [34]. Cells were grown in complete media according to the supplier's recommendations and maintained in a humidified incubator at 37 °C in the presence of 5% CO2. Cells seeded at a density of 4–5 × 103 cells/cm 2 were allowed to grow for 24–48 h and thereafter subjected to real time imaging experiments or immunofluorescence procedures. 2.6.2. Spectroscopic measurements on cells BHK-21 cells were detached from the culture flasks by trypsinization, counted and washed with phosphate buffered saline (PBS) by centrifugation (450 ×g for 5 min) to remove the culture medium. Cells were thereafter incubated (1 min) in 2 mL of PBS containing O1. To remove the unbound dye, loaded cells were quickly centrifuged and washed in PBS. Finally, O1-loaded BHK-21 were resuspended in 2 mL of PBS and subjected to fluorescence measurements in a Varian Eclipse spectrofluorimeter using a 1-cm quartz fluorescence cuvette. The excitation and emission slit widths were 5 nm.

Fluorescence images were captured by a cooled charge-coupled device camera (CoolSNAP HQ, Photometrics, Tucson, AZ) at different time intervals, depending on the specific assay to be performed and corrected for background fluorescence. Z-stacks were acquired on living INS1-E cells with a Nikon Eclipse TE 2000S epifluorescence microscope equipped with a b/w MicroMax 512BFT CCD camera (Princeton Instruments, NJ) using a DeltaRAM monocromator power module (Photon Technology International). The sample was illuminated through a 100× Nikon Fluor oil immersion objective (NA = 1.30) by monochromatic light (at 360 nm). The filter cube used was the same as above. For each field a z-stack was acquired with a z-step of 0.13 μm. Fluorescence images were captured by a cooled CCD camera (CoolSNAP HQ, Photometrics) and corrected for background fluorescence. Image acquisition was performed with the MetaFluor® software (Molecular Devices, MDS Analytical Technologies, Toronto, Canada). Before 3D reconstructions, z-stacks were processed by blind 3D deconvolution with AutoDeblur® 9.1 (AutoQuant software, MediaCybernetics Inc.) with 60 iterations. 2.6.5. Analysis of photobleaching rates of O1 incorporated in living BHK-21 cells Living BHK-21 cells were loaded on the microscope stage with 2.5 μM O1 or 3 μM DPH dissolved in phosphate buffered saline, pH 7.4 for 1 and 5 min respectively. After washing with fresh media, cells were imaged by epifluorescence microscopy while subjected to continuous irradiation at 360 nm with a xenon lamp at 76 mW through the optic fiber attached to the DeltaRAM monocromator power module (Photon Technology International). Before the beginning of the photo-bleaching experiment, the region of interests were drawn on the transmitted light image and the bleaching protocol was started at the very same time of the shutter opening. The fluorescence intensity of at least five cells per coverslip was quantified, averaged and reported as a function of illumination time. The experiments were performed in triplicate for both the dyes.

2.6.3. Fluorescence quenching technique To assess the localization of O1 into the extracellular leaflet of the plasma membrane, a previously described fluorescence quenching technique was used [2]. Briefly, O1 (5 μM) and DHPE-oregon green (2.5 μM) loaded INS1-E cells were perfused on the microscope stage with a Ringer's solution containing 0.5 mg/mL of crystal violet to quench external fluorescence. As expected for markers that selectively stain the outer leaflet of the membrane, the initial fluorescence was quenched by ~99% after ~0.5 min for both the dyes.

2.6.6. Cell viability assay The acute phototoxicity of O1 during 30 min imaging experiments was monitored using the Trypan Blue Exclusion assay [35], a test broadly used to determine the number of viable cells by monitoring the integrity of their plasma membrane. Briefly, BHK-21 cells plated on glass coverslips were loaded on the microscope stage with different O1 concentrations (0 μM, 1.25 μM, 2.5 μM, 5 μM) for 1 min, washed and thereafter subjected to imaging experiments for 30 min, as described in the text. At the end of each experiment, cells were incubated for 5 min on the microscope stage with a Ringer's solution containing 0.2% trypan blue. The viability of the cells was determined by counting the number of nonviable cells (stained blue) present relative to the total number of cells and compared to control sample not loaded with O1 and illuminated in the same experimental conditions. At least five random fields were analyzed for a total of ~ 120 cells/coverslips. The sample counts were performed in triplicate.

2.6.4. Single cell real time experiments Coverslips with attached cells were mounted in an open-topped perfusion chamber (series 20; Warner Instrument Corp, Hamden, CT) on the stage of a Nikon TE2000-U (Nikon Instruments Europe B.V., Bodhoevedorp, The Netherlands). Cells were then loaded at room temperature with 2–10 μM O1 dissolved in extracellular Ringer's solution for 2 min, and thereafter washed in the same buffer. Stained samples were illuminated through a 40 × Nikon Fluor oil immersion objective (NA = 1.30) by monochromatic light (at 360 nm). The emitted fluorescence was passed through a dichroic mirror (410DRLP), and a long pass emitter (435ALP Omega Optical, Brattleboro, VT) classically used for blue emitting fluorophores.

2.6.7. Multicolor labeling of fixed and permeabilized cells 48 h after seeding, BHK-21 cells and INS1-E cells were washed three times in Phosphate Buffered Saline (PBS) at RT and fixed with 4% formaldehyde/PBS for 20 min. After three washes with PBS, cells were permeabilized with 0.1% Triton X-100/PBS. For F-actin visualization, cells were thereafter incubated for 30 mintes with TRITC conjugated phalloidin (10 μg/mL; Fluka), washed three times with PBS, and stained for 10 min at room temperature with O1 (10 μM). After three subsequent washing steps, the coverslips were mounted with 90% glycerol/PBS on glass slides. For insulin granules visualization, fixed and permeabilized INS1-E cells were saturated with 0.1% gelatin in PBS (30 min) and thereafter

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incubated with an anti-insulin specific rabbit antibody (1:100) for 3 h at RT. After three washing steps with 0.1% gelatin in PBS (5 min each), cells were incubated at RT for 30 min with the Alexa Fluor 488 goat anti-rabbit immunoglobulin G secondary antibody conjugate (1:1000 dilution) (Invitrogen). Finally, the coverslips were washed three times with PBS, stained for 10 min at room temperature with O1 (10 μM) and mounted with glycerol/PBS on glass slides. Cells were observed with a 60× oil objective using a LEICA epifluorescence microscope equipped with a NIKON CCD camera, illuminated by a mercury short arc photo optic (HBO 103 W/2 lamp, Osram, Augsburg, Germany). Brightness, contrast adjustments and necessary cropping were performed using the NIH IMAGEJ program.

3. Results and discussion

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For this aim a conjugated backbone comprising a central fluorene and two phenyleneethynylene branches was designed to assure good fluorescent properties. The molecular structure of fluorophores for cell membrane staining requires amphiphilic character to confer balanced solubility in water and in the liphophilic environment of the phospholipid bilayer core. With this aim, 18 carbon atoms alkyl chains were introduced as alkoxy groups on the external phenyl rings, while polar functionalities (sulfonate groups) were attached to the 9 position of the central fluorene ring (compound O1). Such structure is expected to drive the fluorescent and apolar conjugated backbone of the dye to be embedded into the lipophilic core of the cell membrane leaving the sulfonate groups on the external polar surface of the membrane. 3.1. Synthesis of the oligomer O1

With the aim of further expanding the exploration of the applicability of aryleneethynylene-based fluorophores [19,20] in routine cell biology experiments, we elected to design a new membrane marker useful for studies on model membrane systems and classical epifluorescence microscopy approaches. Notwithstanding the previously described (see Introduction) intrinsic weakness of UV-excited aryleneethynylenes as marker for live cell fluorescence microscopy, and the availability of many fluorescent dyes for biological membranes (e.g. lipophilic carbocyanine dyes, fluorescent labeled lectins and phospolipids and the more recently available Cell Mask plasma membrane stains), we were particularly intrigued by the possibility of exploring the use of these quite novel and bright fluorescent cores to obtain low cost, fast and robust membrane staining. In particular, taking advantage of the suitable physico-chemical properties of aryleneethynylenes, such as high yield of fluorescence, optical tunability, easy functionalization and low cost of synthetic procedures, we aimed at tackling a number of every-day bio-chemical applications, such as physico-chemical studies of membrane mimics, and classical epifluorescence microscopy experiments. Finally, the possibility recently proven by others on polymeric compounds based on the aryleneethynylene conjugated backbone [19,20], of utilizing such materials as dyes in two-photon excitation microscopy, prompted us to start this proof-of-concept work on the use of a new weight-defined molecular aryleneethynylene-based marker in studies on liposomes and plasma membranes of mammalian cells.

The synthesis of the compound O1 is based on the palladium-catalyzed cross-coupling reaction [36] between the trimethylsilylethynyl derivative 1 and the aryl iodide 2 in the presence of silver oxide as the base (Scheme 1). The synthesis of the intermediate compounds 1 and 2 is shown in Scheme 2. 3.2. Spectroscopic studies 3.2.1. Spectroscopic properties of O1 in aqueous and DMSO media In order to mimic the hydrophobic environment characteristic of cell membranes and the aqueous medium of extracellular spaces, we started the spectroscopic characterization of O1 using DMSO and HEPES buffer as solvents, respectively. Spectroscopic measurements in DMSO solution showed that O1 is an efficient blue emitter under UV excitation (absorption λmax = 350 nm, εmax 57 × 10 3,emission λmax = 390/410 nm and Φfl = 0.72 relative to quinine sulfate in 0.5 M H2SO4 [37]. Switching from DMSO solution to aqueous medium (20 mM HEPES pH 7.2) the absorption spectrum of O1 showed a small drop in absorbance (Fig. 1A) while the emission spectrum was strongly reduced in intensity (by ~ 5 fold), broader and red-shifted (Fig. 1B) with a maximum at 502 nm. Since one of the most important properties of a good fluorophore for biological applications is its pH stability in physiopathological conditions, we next evaluated O1 sensitivity to extracellular pH changes. Absorption and emission spectra of the marker were thus recorded in

Scheme 1. Synthesis of O1.

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Scheme 2. Synthesis of building blocks 1 and 2.

extracellular Ringer's solutions buffered at different pHs. While a certain pH-dependency of the fluorescence is frequently observed in commonly available probes, the optical properties of O1 are almost independent on pH, in the range 5.0 – 8.0 (Fig. 2). Thus, given its pH stability in the physio-pathological range, even considering the compartments at high H + concentration described in the recent literature in close proximity of the plasma membrane [38,39], O1 represents a good candidate for plasma membrane staining in fluorescence microscopy studies.

3.2.2. Spectroscopic properties of O1 in liposomes Novel organic solutes, such as molecular dyes, represent crucial tools for studies on molecular interactions between micelles/vesicles and biological macromolecules, like mononucleotides and DNA [40,41]. To determine the affinity of O1 for membrane systems and its potential use in liposome-based studies the changes in fluorescence intensity of O1 upon addition of liposomes was investigated. In particular, to gain qualitative insight into the relative affinity of O1 for biological membranes and establish the optimal concentrations for use in cell membrane

Fig. 1. A) Absorption spectra of O1 (4 μM) in Hepes 20 mM (pH 7.2) and DMSO; B) Fluorescence emission spectra of O1 in Hepes 20 mM (pH 7.2) and DMSO.

Fig. 2. Absorption (A) and emission (B) spectra of O1 (1 μM) in extracellular Ringer's solutions buffered at different pH-values. Emission spectra were measured at an excitation wavelength of 360 nm.

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staining experiments, the fluorescence intensity of O1 was monitored as a function of liposome and dye concentrations. When an aqueous solution of O1 (1 μM in 20 mM HEPES, pH 7.2) was titrated with a suspension of cationic liposomes, the dye recovered the spectral features measured in DMSO (λmax =388/410 nm, see the blue trace in Fig. 1B) and showed a significant increase in fluorescence, (Fig. 3). As shown in Figs. 3 (black spectrum) and 1B, in the absence of liposomes the typical peak of the marker in water solution (HEPES) is recordable (500 nm). As the ratio liposome/O1starts to grow, and thus the dye incorporates into membranes (blue spectra), the spectra undergo a blue shift, showing a peak around 420 nm. Since the fluorescence intensity directly reflects the affinity of O1 for the lipidic environment, the data shown in the inset of Fig. 3 give an indication of the relative affinity of the dye towards cationic liposomes [42]. To give a more accurate estimation of the concentrations of dye and liposomes at which the blue shift of the spectrum can be recorded, a titration of 10 μM liposomes with increasing concentrations of O1 (0–2 μM) was performed (data not shown). From this experiment it was established that when the molar ratio liposomes to O1 is greater than 40, the “aqueous fluorophore” concentration can be considered negligible and the classical blue shifted spectrum is recorded. Vice versa, at ratios lower than 40 spectra typical of the “aqueous fluorophore” take place. To gain further insight into the affinity of O1 for lipid membranes, we next performed a titration of a high liposome concentration (100 μM) with low concentrations of O1 (12.5–250 nM) (Fig. 4). As expected, at these molar ratios, all the emission spectra were shifted in the blue region sharing the typical "dye in membrane" peak at 420 nm. While these results highlight the enhancement of O1 fluorescence intensity induced by the presence of the hydrophobic environment, it is clear that an estimation of the enhancing capacity of membranes in respect to O1 optical properties is strongly dependent on the liposomes/dye ratio. All together these data prompted us to test O1 as fluorescent marker for mammalian cell membranes.

3.3. Validation of O1 as membrane marker for microscopic applications in living and fixed mammalian cells 3.3.1. Spectroscopic properties of O1 in cell membranes To directly test if, as suggested by the data collected on liposomes, O1 was able to incorporate within the plasma membrane of mammalian

Fig. 3. Normalized emission fluorescence spectra (excitation 360 nm) during the titration of O1 (1 μM in 20 mM HEPES pH 7.2) with liposomes (1–120 μM). Inset: normalized emission fluorescence intensity at the wavelength characteristics of dye in membrane (410 nm) as a function of liposome concentration.

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Fig. 4. Normalized emission fluorescence spectra (excitation 360 nm) of O1 (12.5– 250 nM) during the titration of 100 μM liposomes.

cells, BHK-21 fibroblasts were loaded with different O1 concentrations and subjected to fluorescence-spectroscopy measurements. In Fig. 5 the excitation and emission spectra of the marker O1 after its anchorage to cell membranes are reported. As expected, the emission spectrum of O1 after its incorporation into cell membranes showed a broad peak shifted toward shorter wavelengths compared with the spectrum in HEPES and a higher similarity to the spectrum of the fluorescence probe O1 dissolved in liposomal membranes (Fig. 3, blue lines).

3.3.2. Fluorescence microscopy studies on living cells To validate the use of O1 as a membrane marker for microscopic applications in living mammalian cells, we marked different cell lines, such as fibroblasts-like cells (BHK-21), insulin secreting beta cells (INS1-E) and primary cultures of neonatal rat cardiac fibroblasts and myocytes. Living cells grown attached to glass coverslips were loaded for 1 min on the stage of an inverted epifluorescence microscope by perfusion with an extracellular Ringer's solution containing different O1 concentrations (2–10 μM) depending on the cell type. The samples were excited at 360 nm and the emitted fluorescence images were collected by a CCD camera and a commercial software for cell real time imaging experiments. The dichroic and emission filters used in these experiments were the same as those routinely used for other known blue emitting fluorescent probes, particularly for Blue Fluorescent Proteins (see Methods for details on the filters and imaging

Fig. 5. Excitation (λem 510) and emission (λexc 360 nm) spectra of O1 (1.25 μM) incorporated in BHK-21 cells. In order to eliminate the contribution of cell-autofluorescence, the excitation and emission spectra of unloaded cells were subtracted from the corresponding spectra of O1-loaded cells.

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Fig. 6. Representative epi-fluorescence images (λexc 360 nm) of living primary cardiac fibroblasts (A) and insulin secreting beta cells (B) loaded on the microscope stage with O1 (10 μM).

setup). In the representative epifluorescence images in Fig. 6A and B, the ability of O1 to highlight the presence of small cell projections (panel a) and cell boundaries (panel b) is apparent. To assess more precisely the membrane localization of O1 in living cells, sequential images of INS1-E cells loaded with O1 were taken at a defined distance (0.13 μm) by means of an epifluorescence microscope equipped with a Z-stack apparatus. A three dimensional reconstruction of the image was achieved by a deconvolution software. Fig. 7 shows a triple view image of two INS1-E cells, in which the membrane labeling is clearly visible. The correct localization of the marker into the extracellular leaflet of the membrane was further assessed by a fluorescence quenching technique [2]. In brief, INS1-E cells loaded with 5 μM O1 were perfused on the microscope stage with a Ringer's solution containing 0.5 mg/mL of crystal violet, a quenching reagent previously used to prove the selective staining of plasma membrane by Evans Blue in living neutrophils.

B

A

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As expected for a dye that selectively stains the plasma membrane, the initial fluorescence of O1 incorporated in INS1E cells was completely quenched (99%) after ~0.5 min. When the same procedure was applied to INS1E cells loaded with a well known plasma membrane marker, DHPE-Oregon Green (Life Technologies), similar results were obtained (data not shown). Another highly desirable feature of fluorochromes to be used in live cell imaging, is the resistance to photo-bleaching. Thus, to validate the suitability of the membrane marker O1 in live-cell real-time imaging experiments, we assessed the bleaching rate of our fluorophore (2.5 μM) in living BHK-21 cells. After loading with O1 BHK-21 cells were continuously illuminated (λexc 360 nm) through the optics of our imaging system (see the Methods section for details). For comparison, parallel experiments were performed with DPH, another UV-absorbing membrane fluorophore, which has been recently employed for liquid droplets staining in Hela cells [43] (Fig. 8). As clearly evidenced by the bleaching rates reported, O1 showed a higher photostability than DPH. In fact, notwithstanding the quite fast bleaching of O1 recordable in the very first seconds of illumination, the fluorescence of O1 remained considerably higher than that of DPH, as measured in the very same experimental conditions in terms of sampling frequency and microscope setup. Since both the sampling frequency and the exposure time are critical parameters that have to be set in each live cell experiment (depending on the biological process in question), we next tested the rate of bleaching of O1 during a typical illumination protocol optimized for studies on membrane dynamics.

10 µm

C

Fig. 7. Triple view image from a z-stack of a group of living INS1-E cells stained with 10 μM of O1. The XY image (panel A) was taken at a middle plane, XZ (panel B) and ZY (panel C) images were taken at the planes indicated by the cross-sections. Image reconstruction was performed with the Autodeblur Software (AutoQuant).

Fig. 8. Photobleaching of O1 loaded on living BHK-21 fibroblasts. Cells were incubated on the microscope stage with 2.5 μM O1 or 3 μM DPH, washed by perfusion with fresh extracellular Ringer's solution and imaged by epifluorescence microscopy while subjected to continuous illumination.

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Setting the dose of illumination (intensity of excitation multiplied by the time of illumination) to the minimum necessary to record clear images and thus minimize both photobleaching and phototoxicity we were able to obtain stable recordings for up to 30 min (see the representative movie in Supplemental Data). Another fundamental parameter to be controlled in live cell imaging experiments is the concentration of the fluorophore. In fact, during the previously described experiments, we observed that at high concentrations of O1 (>5 μM), particularly at high illumination doses, phototoxic phenomena such as membrane blebbing and change of cell shape could occur, independently on any external stimulation. These effects were particularly severe in hormone secreting and excitable cells, such as insulin secreting beta cells (INS1-E) and cardiomyocytes. The observation that cells outside the illuminated area did not show a significant amount of these toxic effects suggested us that the phenomenon can be considered a "phototoxic" effect, most probably due to the short wavelength of the exciting light [21]. The production of toxic radicals (ROS) under these experimental circumstances most probably accounts for such deleterious effects, as it has been described for a number of fluorophores [44]. As already stated in the Introduction, such a mechanism has been indicated as the responsible for the biocidal activity of cationic conjugated polyelectrolytes (CPEs) with backbone based on the phenyleneethynylene repeating unit [23], thus for compounds similar to O1. Importantly, against the idea of a general toxicity of aryleneethynylenes is a recent work specifically focused on the membrane perturbation activity of poly(phenyleneethyneylene) (PPE)based cationic conjugated polyelectrolytes (CPEs) and cationic phenyleneethynylene oligomers (OPEs) [22]. In this paper, the authors clearly showed that only three out of the seven cationic CPEs and OPEs tested against model bacterial and mammalian membranes exerted measurable membrane perturbation against mammalian cell mimics. Moreover, the only anionic CPE tested, PPE-SO3, bearing the sulfonate group (that is the same group of the fluorophore O1) showed no activity against both bacterial and mammalian membrane models.

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Thus we checked if O1 could exert phototoxic effects also in the low-illumination experimental conditions. Importantly, neither BHK-21 cells nor primary cardiac fibroblasts loaded with 1.25–2.5 μM O1 and subjected to "long term" imaging recordings, showed significant membrane blebbing or changes in cell shape (see the movie in Supplemental Data). To further evaluate the phototoxicity of our marker, BHK-21 cells loaded with 1.25 and 2.5 μM O1 were subjected to the trypan blue exclusion assay after the 30 min illumination protocol. As expected by the previous observations, no damage to the cell membranes was measurable (nexp= 3 for each concentration tested). In summary, although the loading conditions and the illumination protocol represent critical experimental factors to be taken into account (as with all the fluorophores used in live-cell imaging) these findings suggest that oligomer O1 can be used as a blue fluorescent probe for real-time imaging of cell membranes in living cells. 3.3.3. Membrane staining of fixed mammalian cells Finally, O1 was tested as a blue fluorescent membrane marker for multicolor labeling in classical immunofluorescence experiments on fixed and permeabilized cells. Notwithstanding the broad emission spectra of the marker and the background signal visible in some experiments, thanks to the brightness and the excitation at short wavelengths of O1, the signal-to-noise ratio was always very good and enabled to obtain clear multicolor images. During the setup of the optimal protocols to be used, it was verified that O1, similarly to most of the available membrane probes, does not resist to the membrane permeabilization step that is necessary for immunofluorescence labeling. Interestingly, and, to our knowledge, differently from most of the plasma membrane markers available, when O1 was used after cell fixation and permeabilization it retained the plasma membrane localization with no significant labeling of intracellular membranes (Fig. 9). In the representative immunofluorescence images depicted in Fig. 9 O1 was used in combination with tetramethylrhodamine phalloidin (TRITC-conjugated phalloidin), a red fluorophore for visualization of

Fig. 9. Multicolor labeling of fixed and permeabilized mammalian cells stained with O1 and red or green fluorophores. (A–C) BHK-21 cells were fixed, permeabilized, and incubated with Phalloidin-TRITC to visualize F-actin (red in panel A). Plasma membranes were then stained with 10 μM of O1. (C) Merge of the two immunofluorescence signals. (D–F) Insulin secretory granules (green in panel D) were visualized in fixed and permeabilized INS1-E cells after immunostaining with an anti‐insulin mouse IgG1 and a green-fluorescent Alexa Fluor®488 goat anti‐mouse IgG.

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F-actin in BHK-21 cells (Fig. 9A–C) or with Alexa Fluor 488, a green fluorophore conjugated to a secondary antibody (see figure legends and Methods for details) for visualization of insulin secretory granules (Fig. 9D–F) in INS1-E cells. Importantly, no significant crosstalk of the blue emission of O1 with the red and green channels was observed by using standard immunofluorescence procedures and optical filter sets (DAPI filters) existing in most fluorescence microscopes. Thus, O1 can be conveniently used in every-day immunofluorescence applications, i.e. as an alternative to the very expensive fluorescent lectins, simply adding it at the end of the antibody incubation procedure and visualizing cells through the optics of a commonly available widefield microscope. As such it can be used, i.e. to measure cell membrane area (such as it is necessary in studies on cardiomyocyte hypertrophy), or visualize small filopodia/lamellipodia in conjunction with other antibodies against different protein of interest. Last, but not less important, during the microscopic study we realized that the use of O1 is quite advantageous in comparison with other commercially available membrane dyes both in term of cost and stability. Most of the available dyes, in facts, are very expensive and highly unstable, so that they must be stored frozen (−20 °C). On the contrary, the new dye herein described is highly stable and can be stored at room temperature as a pure solid or in solution (readily available to use) for up to 4 years. 4. Conclusion In conclusion, we have demonstrated the use of an aryleneethynylene compound as an effective fluorescent marker for liposomes and mammalian cell membranes. It exhibits fast staining (few minutes), insensitivity to pH in the patho-physiological range, good photostability and effective multicolor labeling in immunofluorescence experiments. In summary, the data reported here add substantially to our understanding of the usability of the large class of aryleneethynylene molecules as an entire new family of fluorescent probes for biological studies that will benefit of the broad structural tunability and easy functionalization which has been gained for these emitters in organic optoelectronics applications. Finally, the possibility of using the new dye in two-photon imaging will be the subject of future studies. Supplementary data to this article can be found online at http:// dx.doi.org/10.1016/j.bbamem.2012.06.011. Acknowledgements This work was financially supported by the Ministero dell'Istruzione, dell'Università e della Ricerca (MIUR), “Progetto PRIN 2009 PRAM8L”, and by Fondo per gli Investimenti della Ricerca di Base (FIRB ITA-USA). "grant RBIN04PHZ7". References [1] A.P. Demchenko, Y. Mely, G. Duportail, A.S. Klymchenko, Monitoring biophysical properties of lipid membranes by environment-sensitive fluorescent probes, Biophys. J. 96 (2009) 3461–3470. [2] J. Hed, C. Dahlgren, I. Rundquist, A simple fluorescence technique to stain the plasma-membrane of human-neutrophils, Histochemistry 79 (1983) 105–110. [3] C. Dahlgren, I. Rundqvist, O. Stendahl, K.E. Magnusson, Modulation of polymorphonuclear leukocyte locomotion by synthetic amphiphiles—effect of saturated fatty-acid esters (C2-C18) of poly(ethyleneglycol) 6000, Cell Biophys. 2 (1980) 253–267. [4] R. Ishitsuka, S.B. Sato, T. Kobayashi, Imaging lipid rafts, J. Biochem. 137 (2005) 249–254. [5] L.L. Song, E.J. Hennink, I.T. Young, H.J. Tanke, Photobleaching kinetics of fluorescein in quantitative fluorescence microscopy, Biophys. J. 68 (1995) 2588–2600. [6] R. Sjoback, J. Nygren, M. Kubista, Absorption and fluorescence properties of fluorescein, Spectrochim. Acta, Part A 51 (1995) L7–L21.

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