Surface Nanobubbles Studied by Time-Resolved Fluorescence

Jun 7, 2016 - The impact of surface treatment and modification on surface nanobubble nucleation in water has been addressed by a new combination of ...
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Surface Nanobubbles Studied by Time-Resolved Fluorescence Microscopy Methods Combined with AFM: The Impact of Surface Treatment on Nanobubble Nucleation Nicole Hain, Daniel Wesner, Sergey I. Druzhinin, and Holger Schönherr* Physical Chemistry I, Department of Chemistry and Biology & Research Center of Micro and Nanochemistry and Engineering (Cμ), University of Siegen, Adolf-Reichwein-Strasse 2, 57076 Siegen, Germany S Supporting Information *

ABSTRACT: The impact of surface treatment and modification on surface nanobubble nucleation in water has been addressed by a new combination of fluorescence lifetime imaging microscopy (FLIM) and atomic force microscopy (AFM). In this study, rhodamine 6G (Rh6G)-labeled surface nanobubbles nucleated by the ethanol−water exchange were studied on differently cleaned borosilicate glass, silanized glass as well as self-assembled monolayers on transparent gold by combined AFM-FLIM. While the AFM data confirmed earlier reports on surface nanobubble nucleation, size, and apparent contact angles in dependence of the underlying substrate, the colocalization of these elevated features with highly fluorescent features observed in confocal intensity images added new information. By analyzing the characteristic contributions to the excited state lifetime of Rh6G in decay curves obtained from time-correlated single photon counting (TCSPC) experiments, the characteristic short-lived (1 μm in the surface normal direction).36,37 Ohl and co-workers have pioneered the use of total-internal-reflection-fluorescence (TIRF) microscopy to observe dye-labeled surface nanobubbles38 and showed that alleged surface nanobubbles and small particles can be successfully differentiated in microfluidic experiments.39 A very recent study introduced a new technique based on TIRF and photoactivatable dyes to analyze interfaces supporting nanobubbles.40 An alternative approach reported by Karpitschka et al. analyzed the nucleation of surface nanobubbles on glass by label-free optical interference-enhanced reflection microscopy and showed colocalization of alleged surface nanobubbles by optical and atomic force microscopy.41 Finally, the change in fluorescence properties of a surface immobilized dye was shown to allow one to localize nanobubbles.42 We have recently developed a novel high-resolution optical approach to confirm the gaseous nature of surface nanobubbles and to differentiate them from known contaminants, among others.43 By exploiting the approach by Ohl et al. to label the alleged air−water interface of surface nanobubbles with a suitable fluorescent dye,38 time-resolved fluorescence microscopy methods are used in this approach to differentiate local nanoenvironments of the reporter dyes (Scheme 1). By combining the so-called fluorescence lifetime imaging microscopy (FLIM) approach with AFM in one setup, individual surface nanobubbles can be studied by two

complementary techniques, and the new optical data hence obtained can be related directly to AFM observations reported earlier. As we report in this Article, this combined AFM-FLIM approach can be applied as powerful combination to elucidate the role of the substrate in nanobubble nucleation, to analyze the shape and to confirm the gaseous nature of surface nanobubbles on very differently functionalized substrates. In particular, surface nanobubbles nucleated by the ethanol−water exchange on differently cleaned glass, silanized glass exposing methyl and amino groups and thiol-modified gold layer were comparatively analyzed.

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EXPERIMENTAL SECTION

Preparation of the Dye Solutions. Rhodamine 6G, Rh6G (purity ∼95%, Fluka, Germany) was used as fluorescent dye with a concentration of 850 nM and was either dissolved in Milli-Q-water (obtained from a Millipore Direct Q 8 system (Millipore, Schwalbach, Germany) with resistivity of 18.2 MΩ cm or in ethanol (EMSURE ACS, 99.9%, ISO, Reag. PhEur., Merck Millipore, Darmstadt, Germany). Preparation of the Surfaces. Glass coverslips (20 mm × 20 mm with a thickness of 0.13 to 0.16 mm (borosilicate Menzel-Gläser, Gerhard Menzel GmbH, Braunschweig, Germany) were cleaned first with chloroform (ROTIPURAN ≥ 99% p.a., ROTH, Karlsruhe, Germany), followed by rinsing with ethanol and then Milli-Q water. Afterward, the substrates were placed in piranha solution (a 3:1 v/v mixture of sulfuric acid (CHEMSOLUTE, 95%, p.a, Rennigen, Germany) and hydrogen peroxide (30%, Ph. Eur, ROTH, Karlsruhe, Germany)) for 10 min, and rinsed with water and then with pure ethanol. (Caution! Piranha solution is very reactive and may explode upon contact with organic materials and solvents. Extreme precautions must be taken at all times.) For the modification of the glass slides with 1-octadecanthiol (ODT; HS-(CH2)17CH3, 98%, purchased from Aldrich), the piranha cleaned glass slides were placed in the vacuum chamber of a thermal evaporator (MED 010, Balzers Union, Liechtenstein) for evaporation of a 2−5 nm thin layer of titanium (at a pressure of 1 × 10−6 mbar with an evaporation rate of ∼0.1 nm/s). On top of the titanium, a transparent layer of gold was evaporated with a thickness between 12 to 15 nm. The evaporation of titanium and gold was done in one evaporation run. After the evaporation of the gold, the glass slides were allowed to cool down for 30 min and then placed in a 1 mM ethanolic 11156

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channel. The reference fluorescence decays of Rh6G in aqueous solutions and at the glass−water and at water−air interfaces, respectively, were obtained from the time correlated single photon counting (TCSPC) representation of TTTR data. To determine the fluorescence lifetime at the water−air interface, a drop of aqueous Rh6G solution was placed on the bottom of a glass coverslip and the focus of an air objective with 40× magnification (Plan N 40x/0.65, Olympus, Hamburg, Germany) was moved to the interface. For the degassing experiments, a 250 mL aqueous R6G solution was degassed for 3 h at 20 °C by using a diaphragm vacuum pump (Type MZ 2C, Vacuubrand, Germany). The pressure was set to 10 mbar, while the solution was stirred continuously. Combined AFM-FLIM Measurements. The AFM images of surface nanobubbles were acquired under liquid with a resolution of 512 × 512 pixels2 with a scan rate of 0.5 Hz, an amplitude of 50 nm and an amplitude set point ratio around 87−90%, respectively. The samples were prepared on glass cover slides mounted in the Closed Fluid Cell (Asylum Research). This cell is airtight, but allows for the solvent exchange via several ports by an external pump, while the AFM tip stays close to the surface. The cantilever was cleaned prior to the measurements in an UV-Ozone cleaner (ProCleaner, UVOTECH Systems, Inc., Walnut Creek, CA). When the nanobubbles were nucleated with glass Pasteur pipettes, the ethanol−water exchange was performed in the AFM liquid cell and then afterward the AFM-head with the cantilever holder was mounted. For experiments with degassed water, the degassed aqueous Rh6G solution was pumped through the AFM liquid cell (see Supporting Information for all further details). In the combined AFM-FLIM setup the tip of the AFM has to be first aligned with the laser focus of the optical microscope by using the AFM top-view camera and either the beam diagnostic camera in the main optical unit or the backscattering of the laser focused onto the tip to monitor the position of the AFM probe with respect to the laser beam. AFM and FLIM data were acquired sequentially using the separate control units to avoid interferences. Due to the practically static situation (on experimental time scales) following nanobubbles nucleation, this did not affect the data. Image Processing. For the analysis of the acquired fluorescence intensity images, in particular the cross section analyses, ImageJ1.47v software (from Wayne Rasband, National Institutes of Health, USA) was used. For the overlay of the intensity images in section 4.3.2, a conversion of the data into binary images with a certain threshold for high and low intensity pixel was carried out. The overlay and further analysis of intensity maps was done and analyzed with MATLAB (release R2010a, MathWorks, Natick, MA) software. AFM images were flattened using first-order polynomials and masking the elevated structures of the nanobubbles utilizing the implemented software (Asylum Research) or SPIP (Image Metrology, Horsholm, Denmark). Fitting Procedure for Determination of the Fluorescence Lifetimes. The parameters for the fluoresce decay of Rh6G in a water and at the different interfaces were obtained by employing a homemade software, based on a nonlinear gradient least-squares method,46 with the global sum of the weighted squares of the residuals

solution of ODT for 2 h, then rinsed of with ethanol and blown dry in a steam of nitrogen. Glass slides were also modified with 3-aminopropyl(dimethylethoxysilane) (APMES, Aldrich, 97%) or octadecyltrichlorosilane (OTS, Arcos Organics, 95%).44,45 For this purpose the freshly cleaned glass slides were placed in a desiccator, in which a glass beaker with a small amount of the silane (OTS, APMES) was placed. The desiccator was then evacuated with a pump with 0.4 mbar for 1 h and was kept under vacuum for 12 h at 20 °C. For the experiment where the effect of different glass treatments on the surface nanobubbles was evaluated, the glass cover slides were first rinsed with ethanol and blown dry with a steam of nitrogen. Then they were placed in the UV-Ozone cleaner (ProCleaner, UVOTECH Systems, Inc., Walnut Creek, CA) for 15 min or cleaned by oxygen plasma (Plasma Prep II, SPI Supplies, West Chester, USA) for 10 min. Contact Angle Measurements. Static contact angle measurements of 2 μL Milli-Q water were performed with minimal delay after the preparation of the surfaces with an OCA 15plus instrument (DataPhysics Instruments GmbH, Filderstadt, Germany). Surface Nanobubble Nucleation. The ethanol−water exchange was performed with two Eppendorf pipettes (Eppendorf Research plus, 100 μL-1000 μL, Hamburg, Germany) or two piranha-cleaned glass Pasteur pipettes to nucleate surface nanobubbles. Initially, 400 μL of the aqueous dye solution (850 nM) was placed on a clean coverslip mounted in the confocal cell and then imaged at the desired position. Two pipettes were then used to simultaneously pipet in and out liquid volume of 400 μL for the exchange. In a first step, the aqueous dye solution was exchanged by an ethanolic dye solution (850 nM), by pipetting out the former and simultaneously pipetting in the latter solution. This process with new portion of dye solution was repeated three times to ensure that practically all water was exchanged to ethanol. In a second step, the actual ethanol-water exchange, the ethanolic dye solution was pipetted out of the confocal cell and aqueous dye solution (850 nM) was simultaneously pipetted in. This procedure was repeated three times as well. AFM Measurements. The AFM images of surface nanobubbles were taken using intermittent contact (tapping) mode done on a MFP-3D-Bio atomic force microscope (Asylum Research, Santa Barbara, CA) using for V-shaped MLTC Si3N4 cantilevers (Bruker AXS, Camarillo, CA) with a nominal resonance frequency of 120 kHz and a nominal spring constant of 0.6 N/m. The data were acquired with 512 × 512 pixel2 with a scan rate of 0.5 Hz and an amplitude of 43 nm and amplitude set point ratio around 86−92%, respectively. The samples were prepared on glass cover slides mounted in the Closed Fluid Cell (Asylum Research) for measurements in liquid. The AFM roughness measurements of the differently treated glass surfaces were performed in air in intermittent contact mode, with AC160 cantilevers (Olympus, Tokyo, Japan), with amplitude of 1.0 V and a set point around 90−95%, respectively. A scan area of 1 μm × 1 μm scanned with a resolution of 512 × 512 pixels2. FLIM Measurements. Fluorescence lifetime imaging was performed using an inverted microscope (IX 71, Olympus, Hamburg, Germany), a commercial main optical unit (Microtime 200, PicoQuant, Berlin, Germany), a fiber coupling unit (FCU II, PicoQuant) and a data acquisition module (PicoHarp 300, PicoQuant). The sample was excited by a pulsed diode laser (LDHD-C-485, PicoQuant) at 485 nm with a fwhm of 55 ps and repetition rate of 20 MHz. The laser power was attenuated with neutral density filters. The excitation light was focused on the sample and the fluorescence emission was collected by a water-immersion objective with 60× magnification (UPlanSApo 60x/1.20 N.A., Olympus, Hamburg, Germany). The fluorescence was detected by a SinglePhoton Avalanche Diode (PD1CTC, Micro Photon Devices, Bolzano, Italy) after passage through a dichroic mirror (z500dcxr, Chroma, Bellows Falls, USA), and an emission filter (HQ510lp, Chroma, Bellows Falls, USA), and a 50 μm pinhole. The images (512 × 512 pixels2) were recorded with at least 12 000 time bins per pixel, and the time per pixel was 0.6 ms. The measurements, which were done at a temperature of 22 °C, were carried out in time-tagged time-resolved (TTTR) data acquisition mode with a time resolution of 16 ps/

S=

∑ ∑ wi(yij − yije )2 j

(1)

i

as the target function. In the expression for S, w is the weighting factor of the response function

yj (t ) =

∫0

t

⎛ ⎛ t ⎞⎞ p(t − x)⎜⎜∑ ajk exp⎜ − ⎟⎟⎟ dx ⎝ τk ⎠⎠ ⎝ k

(2)

calculated as a convolution of the excitation pulse p and the sum of 1 to 3 exponents. In such a global fit, the decay times variables τn are kept the same for all response functions. The index j, usually 4 to 9, refers to the response functions, and i counts the experimental points for each response function and superscript e indicates that the quantity is an experimental value. For the fluorescence decay, the factor w equals y−1. The response functions y employed here are the first 40 ns of the measured Rh6G fluorescence decays. In decay curve figures, the 11157

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Figure 1. Constant amplitude AFM height images captured in intermittent contact mode at the glass−liquid interface. The data shown in panel a were acquired in high purity water, which was subsequently exchanged to ethanol (panel b) and back to water (panel c).

Figure 2. Confocal fluorescence microscopy intensity images of: (a) the glass/water interface (850 nM aqueous Rh6G), (b) the glass/ethanol interface after exchanging the aqueous medium with 850 nM ethanolic Rh6G, and (c) the glass/water interface after exchanging the ethanolic solution with 850 nM aqueous Rh6G. amplitudes of the acquired lifetimes were normalized to the highest values.

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agreement with the work by the groups of Zhang and Lohse.7,16,21,34 However, unlike following this indirect degassing strategy, which cannot easily differentiate a volatile water-insoluble organic liquid from gas, the work discussed here exploits FLIM, which identifies the gas-filled surface nanobubbles via the characteristic emission component lifetime of the reporter dye Rh6G, when it is partitioned to the gas−water interface. We have verified in independent measurements that water surface tension demonstrates some decrease in the presence of Rh6G. It seems to be an adequate dye for this purpose, as Rh6G is enriched at the air−water interface, hence resulting in a bright fluorescence compared to the solid−water interface as well as the bulk water phase. In the confocal fluorescence microscopy data shown in Figure 2, the focal plane of the microscope was adjusted such that the solid−water interface was residing in the voxel analyzed. While the emission of Rh6G, when excited at 485 nm, is distributed evenly in Figure 2a,b, there is a markedly changed emission intensity image seen in Figure 2c. Highly fluorescent circular features appear in high number density after the ethanol−water exchange, which is known in the literature and has been independently shown above to nucleate alleged surface nanobubbles (Figure 1). The observation of bright fluorescent, i.e., Rh6G labeled features in the confocal intensity image agrees well with the TIRF data by the Ohl group.38 Despite the fact that a fraction of the dye dissolved in water must be excited due to the dimensions of the confocal voxel (in this particular case: Δx = Δy = 150 nm, Δz = 410 nm), the local enrichment at the presumed gas−water interface results in the clear localization. To couple the established AFM observations (e.g., Figure 1) and the new approach (Figure 2), the corresponding experiment was then carried out quasi-simultaneously. As is

RESULTS AND DISCUSSION

To address the potential impact of the surface treatment of the borosilicate glass substrates and functionalization with silane layers and gold supporting a thiol-based self-assembled monolayer, surface nanobubble nucleation by the ethanol− water exchange was first analyzed by conventional AFM, followed by confocal fluorescence microscopy experiments with Rh6G labeled samples and finally the combined AFM-FLIM setup. In situ AFM height data acquired in intermittent contact (tapping) mode in water on piranha-cleaned glass unveiled the presence of some elevated features resembling nanobubbles and potentially some contamination even prior to the ethanol− water exchange (Figure 1a). The formation of surface nanobubbles by gas entrainment is known in the literature, which can explain those elevations.47 After exchanging the water with ethanol and equilibrating the microscopy system, only very few elevated features remained on the glass (Figure 1b). By contrast, after exchanging the ethanol again with water, the surface was found to be covered with round structures that correspond to previous work reporting on surface nanobubbles (Figure 1c). Even though no disposable syringes or needles, which are known to introduce polysiloxane based contamination into the liquid cell and onto the sample surface,21,43 were used in our experiments, there is further evidence required to unequivocally assign those features indeed to gas-filled surface nanobubbles. In our laboratories, we have repeatedly confirmed that similar features vanish, if the cell is purged with degassed water, in full 11158

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Figure 3. Microscopy data captured on surface nanobubbles nucleated by an ethanol−water exchange on OTS/glass: (a) Confocal fluorescence microscopy intensity image; (b) Constant amplitude AFM height image acquired practically simultaneously in intermittent contact mode in the same area as the data in panel a; (c) Overlay of the fluorescence intensity image and the AFM height image.

Figure 4. Microscopy data captured on surface nanobubbles nucleated by an ethanol−water exchange on piranha-cleaned glass in an aqueous 850 nM Rh6G solution: (a) Constant amplitude AFM height image obtained in intermittent contact mode; (b) Confocal fluorescence microscopy intensity image acquired practically simultaneously in the same area as the data in panel a; (c) Triple exponential TCSPC fluorescence decay curve of Rh6G obtained from TTTR data of panel b. The decay times τ and corresponding relative amplitudes a (see eq 2) are shown. The weighted deviations σ (see eq 1), the autocorrelation functions A−C, and the values for χ2 are also indicated.

systems this fluorescence lifetime often strongly depends on the local environment, temperature, excited state quenching and photoinduced chemical reactions. For instance, the fluorescence lifetime of 4-(dimethylamino)benzonitrile was shown to change from 3800 ps in n-hexane to 4.0 ps (the main component of the double exponential decay) in acetonitrile at 25 °C and from 4120 ps in 2-methylpentane at −151 °C to 163 ps in n-hexadecane at 284 °C, i.e., more than 1000 times.48,49 Fluorescence decay times also provide a handle to the study of interfaces. For instance, the influence of a dielectric interface on the fluorescence decay time of a fluorophore was shown by Drexhage.50 For inhomogeneous systems, such as nanobubbles at the liquid−solid interface, the use of a proper dye labeling the interfaces between liquid, solid, and gas hence may help to map the decay component amplitudes ak (see eq 2) with adequate lateral resolution and to identify the nature of the interfaces. In this case the number of fluorescence decay components (n) should be equal to the dimension of the system, the number of interfaces (m) plus one corresponding to the aqueous dye solution.

shown in Figure 3, the elevated features observed only after the ethanol−water exchange on an OTS hydrophobized glass coincide very well with the highly fluorescent patches in the fluorescence intensity image after correcting for a minor offset that is due to limited accuracy of the tip−focus alignment. Since the offset in the xy plane affects all fluorescence features in the same manner, it can be concluded that there is very good colocalization of surface nanobubbles (AFM) and the bright fluorescent objects. The fuzzy fluorescence image with bright features of approximately identical size (diameter ≈ 500 nm) agrees with an image of an ensemble of objects that are similar or smaller in size than the diffraction limit of the confocal microscopy objects. Compared to this limited lateral resolution, the AFM affords much higher resolution data and the well-known metrology within the limitations pointed out in previous reports.27−30 The full potential of the combined confocal-AFM setup is unleashed, when the fluorescence data is not merely displayed as an intensity map, but when time-resolved analyses are carried out. In the setup used, not only the overall amount of emitted photons Fij, but also fluorescence kinetics f ij(t) for each ij-pixel are detected. Fij =

∫0

m=n−1

In the present work Rh6G, which is a well-established tracer dye, was employed for the labeling the nanobubble interfaces. Its photophysics has been studied in different media51 and at different interfaces,52−54 and it is known that the fluorescence lifetime is influenced by concentration and aggregation of the Rh6G.55 We have independently shown that the fluorescence

∞

fij (t ) dx

(4)

(3)

Usually the fluorescence decay (2) of the dye can be described by a single exponential, k = 1. Even in homogeneous 11159

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Figure 5. Zoom of the microscopy data shown in Figure 4: (a) Constant amplitude AFM height image obtained in intermittent contact mode; (b) Confocal fluorescence intensity image of the area shown in panel a; (c) Overlay of both images of panels a and b.

previously established characteristic decay component of Rh6G at the air−water interface. This short component is tentatively attributed to a shortened Rh6G fluorescence decay due to the surface enrichment of Rh6G at the air−water interface,43 known from our surface tension measurements, as well as to the excitation light scattering at this interface. Currently, we do not have solid, i.e., spectroscopic evidence for the possible occurrence of a distinct lifetime of Rh6G at this interface that is indeed very short. Hence, the bright features in the confocal fluorescence images, which possess the short-lived component in the fluorescence decay, and the features observed by AFM can indeed be assigned to gaseous surface nanobubbles. In case of pinning, the contact angle values of surface nanobubbles were reported to be solely a function of gas concentration in the water. The model put forward by Zhang and Lohse7 that relies on contact line pinning explains surface nanobubble stability by the balance of gas outflux (due to the Laplace pressure) and the influx due to oversaturation of the liquid with gas. If the Young equation was valid, surface nanobubbles should not exist on glass. Since surface nanobubbles may however be pinned, the Young equation is not applicable to estimate meaningful values for the contact angle. Systematically acquired data on comparably textured or rough surfaces could support the universality of the contact angle values (at constant gas saturation level) proposed by the abovementioned model.56 Most data discussed in the literature thus far suffer from the fact that different types of solid substrates differ in terms of roughness, which renders the data difficult to compare. The data on unmodified glass discussed above show that nanobubbles were nucleated by the ethanol−water exchange on piranha cleaned glass. The nucleation of surface nanobubbles on similarly cleaned glass has also been reported by several other groups.41 The strongly oxidizing piranha treatment, which is widely used in the field of surface science to clean various solid substrates due to the chemical etching of contaminants and also the surface by the strongly oxidizing solution, affords completely wettable glass surfaces that exhibit static water contact angles OTS (θ = 98°) > ODT (θ = 110°). By contrast, no correlation with the

of energetic plasma particles created from oxygen. Although the UV energy is effective in breaking most of the organic bonds of surface contaminations, UV-ozone cleans the surface mainly with ozone. One may speculate that UV-ozone and plasma cleaning are the milder to the surface, while piranha cleaning is known to etch the surface and to most likely increase the roughness of the surface. After carrying out the exact same ethanol−water exchange protocols elucidated above on glass that was subjected to UVozone and oxygen plasma treatments, no nanobubbles could be observed by confocal fluorescence, FLIM and combined AFMFLIM experiments (Figures 6 and S-2). The double exponential decay for these systems (Figures 6c and S-2c) observed here indicates in accordance with eq 4 that m = 1, i.e., only one water−glass interface is populated with the dye. The fluorescence intensity images (Figures 6b and S-2b) showed homogeneous fluorescence emission of the reporter dye Rh6G at these interfaces, while the AFM scans did not unveil any elevated nanoscale feature (Figure 6a and S-2a). The two decay times observed in Figure 6c, which are similar to those of the long-lived components in Figure 4c, as well as to the decay times of Rh6G in water and on the water−glass interface,43 respectively, do not indicate the presence of an air−water interface in the vicinity of the glass surface. Hence the absence of any short-lived component in the fluorescence decay curves underlines that there are no nanobubbles on these glass surfaces. A plausible explanation is the absence of pinning sites, required according to refs 7, 22, and 23 for the nanobubbles nucleation, on the glass cleaned by the milder UV-ozone and oxygen plasma procedures might be associated with isolated nanometer-scale corrugations rather than with different chemical composition of the surface. The roughness values of those surfaces, however, did not differ markedly. While the UVozone cleaned glass had a roughness (measured on an area of 1 11161

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ACKNOWLEDGMENTS

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REFERENCES

The authors gratefully acknowledge Dipl.-Ing. Gregor Schulte (Physical Chemistry I, Department of Chemistry and Biology, University of Siegen) for advice and excellent technical support, and gratefully acknowledge financial support from the Deutsche Forschungsgemeinschaft (DFG Grant No. INST 221/87-1 FUGG), the European Research Council (ERC grant to H.S., ERC Grant Agreement No. 279202), and the University of Siegen.

(1) Parker, J. L.; Claesson, P. M.; Attard, P. Bubbles, cavities and the long-ranged attraction between hydrophobic surfaces. J. Phys. Chem. 1994, 98, 8468−8490. (2) Lou, S. T.; Ouyang, Z. Q.; Zhang, Y.; Li, X. J.; Hu, J.; Li, M. Q.; Yang, F. J. Nanobubbles on solid surface imaged by atomic force microscopy. J. Vac. Sci. Technol., B: Microelectron. Process. Phenom. 2000, 18, 2573−2575. (3) Ishida, N.; Inoue, T.; Miyahara, M.; Higashitani, K. Nano bubbles on a hydrophobic surface in water observed by tapping-mode atomic force microscopy. Langmuir 2000, 16, 6377−6380. (4) Tyrrell, J. W.; Attard, P. Images of nanobubbles on hydrophobic surfaces and their interactions. Phys. Rev. Lett. 2001, 87, 176104. (5) An, H. J.; Liu, G. M.; Atkin, R.; Craig, V. S. J. Surface nanobubbles in nonaqueous media: Looking for nanobubbles in DMSO, formamide, propylene carbonate, ethylammonium nitrate, and propylammonium nitrate. ACS Nano 2015, 9, 7596−7607. (6) Borkent, B. M.; de Beer, S.; Mugele, F.; Lohse, D. On the shape of surface nanobubbles. Langmuir 2010, 26, 260−268. (7) Lohse, D.; Zhang, X. H. Pinning and gas oversaturation imply stable single surface nanobubbles. Phys. Rev. E 2015, 91, 031003. (8) Peng, H.; Birkett, G. R.; Nguyen, A. V. Progress on the surface nanobubble story: What is in the bubble? Why does it exist? Adv. Colloid Interface Sci. 2015, 222, 573−580. (9) An, H. J.; Liu, G. M.; Craig, V. S. J. Wetting of nanophases: Nanobubbles, nanodroplets and micropancakes on hydrophobic surfaces. Adv. Colloid Interface Sci. 2015, 222, 9−17. (10) Stocco, A.; Möhwald, H. The Influence of Long-Range Surface Forces on the Contact Angle of Nanometric Droplets and Bubbles. Langmuir 2015, 31, 11835−11841. (11) Mishchuk, N.; Ralston, J.; Fornasiero, D. Influence of very Small Bubbles on particle/bubble Heterocoagulation. J. Colloid Interface Sci. 2006, 301, 168−175. (12) Wu, Z. H.; Zhang, X. H.; Zhang, X. D.; Sun, J. L.; Dong, Y. M.; Hu, J. In situ AFM observation of BSA adsorption on HOPG with nanobubble. Chin. Sci. Bull. 2007, 52, 1913−1919. (13) Liu, G. M.; Wu, Z. H.; Craig, V. S. J. Cleaning of Protein-Coated Surfaces Using Nanobubbles - An Investigation Using a Quartz Crystal Microbalance. J. Phys. Chem. C 2008, 112, 16748−16753. (14) Yang, S.; Duisterwinkel, A. Removal of Nanoparticles from Plain and Patterned Surfaces Using Nanobubbles. Langmuir 2011, 27, 11430−11435. (15) Liu, G. M.; Craig, V. S. J. Improved Cleaning of Hydrophilic Protein-Coated Surfaces using the Combination of Nanobubbles and SDS. ACS Appl. Mater. Interfaces 2009, 1, 481−487. (16) Lohse, D.; Zhang, X. H. Surface nanobubbles and nanodroplets. Rev. Mod. Phys. 2015, 87, 981−1035. (17) Craig, V. S. J. Very small bubbles at surfaces-the nanobubble puzzle. Soft Matter 2011, 7, 40−48. (18) Langmuir 2016, 32 (issue), special issue on “Nanobubbles”. (19) Berkelaar, R. P.; Seddon, J. R. T.; Zandvliet, H. J. W.; Lohse, D. Temperature Dependence of Surface Nanobubbles. ChemPhysChem 2012, 13, 2213−2217. (20) Schönherr, H.; Hain, N.; Walczyk, W.; Wesner, D.; Druzhinin, S. I. Surface Nanobubbles Studied by AFM Techniques - Facts, Fiction and Open Questions. Jpn. J. Appl. Phys. 2016, in press.

Figure 8. Histogram of fractional area occupied by surface nanobubbles after ethanol−water exchange under similar conditions, as observed in Figures 2 and 7. On UV-ozone and oxygen plasmatreated glass, no nanobubbles were detected.

roughness data, as analyzed by AFM, was found (see Figure S-5 and Table S-1, Supporting Information).

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CONCLUSIONS The combination of fluorescence lifetime imaging and atomic force microscopy revealed that rhodamine 6G-labeled surface nanobubbles nucleated by the ethanol−water exchange indeed contain gas. The number density of the surface nanobubbles was found to decrease with increasing hydrophobicity on silanized glass and hydrophobic self-assembled monolayers on transparent gold. With the rapid screening option of confocal microscopy imaging, backed by the time-resolved data in FLIM and complementary 3D topography information from quasi simultaneously captured AFM data, it was established that UVozone or oxygen plasma cleaned glass did not support nanobubbles after the ethanol−water exchange in contrast to piranha-cleaned glass. While the origin of this observation could not be conclusively unveiled yet, the potential of the combined AFM-FLIM approach in systematic studies of individual surface nanobubbles and related interfacial phenomena has been confirmed.

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ASSOCIATED CONTENT

* Supporting Information S

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.langmuir.6b01662. Detailed description of the degassing procedure and experiments; statistical analysis of AFM data on surface nanobubbles; AFM, confocal microscopy data, and decay curve for oxygen plasma cleaned glass, XPS survey spectra of differently cleaned glass substrates; analysis of fluorescence intensity images; AFM height data of substrates; and associated roughness data (PDF)

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AUTHOR INFORMATION

Corresponding Author

*Tel.: +49 271 740 2806. Fax: +49 271 740 2805. E-mail: [email protected]. Notes

The authors declare no competing financial interest. 11162

DOI: 10.1021/acs.langmuir.6b01662 Langmuir 2016, 32, 11155−11163

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DOI: 10.1021/acs.langmuir.6b01662 Langmuir 2016, 32, 11155−11163