Letter pubs.acs.org/NanoLett
Cathodoluminescence-Activated Nanoimaging: Noninvasive NearField Optical Microscopy in an Electron Microscope Connor G. Bischak,† Craig L. Hetherington,†,§ Zhe Wang,⊥ Jake T. Precht,† David M. Kaz,†,§,○ Darrell G. Schlom,⊥,¶ and Naomi S. Ginsberg*,†,‡,§,∥,∇ †
Department of Chemistry and ‡Department of Physics, University of California, Berkeley, California 94720, United States Physical Biosciences and ∥Materials Sciences Divisions, Lawrence Berkeley National Laboratory, Berkeley, California 94720, United States ⊥ Department of Materials Science and Engineering, Cornell University, Ithaca, New York 14853, United States ¶ Kavli Institute at Cornell for Nanoscale Science, Cornell University, Ithaca, New York 14853, United States ∇ Kavli Energy NanoSciences Institute, University of California, Berkeley, California 94720, United States §
S Supporting Information *
ABSTRACT: We demonstrate a new nanoimaging platform in which optical excitations generated by a low-energy electron beam in an ultrathin scintillator are used as a noninvasive, near-field optical scanning probe of an underlying sample. We obtain optical images of Al nanostructures with 46 nm resolution and validate the noninvasiveness of this approach by imaging a conjugated polymer film otherwise incompatible with electron microscopy due to electron-induced damage. The high resolution, speed, and noninvasiveness of this “cathodoluminescence-activated” platform also show promise for super-resolution bioimaging. KEYWORDS: cathodoluminescence, nanoimaging, nanostructures, soft materials, super-resolution imaging, resonant near-field coupling
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materials19 and more recently to characterize a variety of metallic nanostructures.20−24 New CL approaches have enabled both mapping directional emission with angular-resolved detection,25 spatially resolving carrier transport through integration of electron and near-field optics,26,27 and hyperspectral imaging on the nanometer scale.28 CL has been used to image biological samples. Yet, direct CL of stained dehydrated samples29,30 did not hold up well to electron damage, and although inorganic cathodoluminescent nanoparticle labels31−33 are more robust, imaging with nanoparticle labels remains invasive because the electron beam must penetrate into the sample, precluding repeated measurements or observations of dynamics. By contrast, to take advantage of the tight focus of an electron beam for spectrally specific and noninvasive imaging, our aim is to use optical excitations generated by a nanoscale electron beam in a cathodoluminescent material above the sample as a noninvasive, near-field optical scanning probe. Although some efforts have been exploring using quantum dot films34 or moderately cathodoluminescent materials35 to generate hybrid electron and optical scanning probes, we recently proposed to combine the nanoscale focus of electron
he emergence of far-field super-resolution techniques, such as stochastic localization,1−4 stimulated emission depletion (STED),5 and structured illumination6 microscopies has revolutionized the fluorescence imaging of labeled biological structures. Yet, capturing nanoscale biological dynamics and imaging systems using their endogenous chromophores both remain challenging for these methods. Near-field optical probes7−17 have proved valuable for the characterization of complex structures and of processes in solid state materials, soft matter, and biological samples that occur over length scales smaller than the wavelength of light. In most variants of nearfield scanning optical microscopy (NSOM), an optical probe is integrated with a scanning tip and rastered over a sample to form an image. Yet, images acquired with NSOM require mechanical scanning and can contain artifacts from tip−sample interactions. On the other hand, in scanning electron microscopy (SEM), a focused electron beam is electronically scanned over a sample to obtain nanoscale images by correlating the detected scattered electrons with the position of the beam, recently achieving the resolution to image single atoms.18 Traditional electron microscopy is incapable of spectrally specific excitation and damages soft materials such as biological samples. One can, however, detect light generated in the sample by the electron beam in a process called cathodoluminescence (CL), which has been used historically to investigate the nanoscale properties of solid luminescent © XXXX American Chemical Society
Received: February 20, 2015 Revised: April 2, 2015
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DOI: 10.1021/acs.nanolett.5b00716 Nano Lett. XXXX, XXX, XXX−XXX
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Figure 1. (a) Imaging chips consisting of a YAlO3:Ce scintillator film supported by LaAlO3 and SrTiO3 buffer layers and a Si frame. Al nanostructures embedded in SiO2 are positioned below and directly against the scintillator film. The ProTEK B3 layer serves as a protective layer during the Si wet etch. (b) Secondary electron (SE) image of the imaging device showing four separate “windows” for imaging. (c) Far-field radiation emitted from the scintillator film is collected by a parabolic mirror positioned above the imaging chip and is then directed outside the scanning electron microscope (SEM) to a photomultiplier tube (PMT) or spectrometer. (d) Close-up of the extent of electron scattering above a SiO2/Al boundary demonstrates that the electrons do not interact directly with the Al, but excite the Ce3+ dopants in the YAP:Ce layer within the volume delineated by the red dotted line. Atomic force microscopy (AFM) (e) and SE (f) images of the Al nanostructures deposited on YAP:Ce prior to encapsulation in SiO2 show that they are 50 nm thick with a point-to-edge distance of approximately 270 nm.
To create a nanoscale excitation spot in a YAP:Ce film, a focused, low-energy (1−2 keV) electron beam in a SEM scatters within the YAP:Ce film and locally excites the film’s Ce3+ dopants (Figure 1a). To use the scintillator film as an excitation source for noninvasive near-field nanoimaging, the focused electron beam must access the film from one side while the sample is positioned directly adjacent to the scintillator film on the opposite side. As the result of our fabrication process, as shown in Supporting Information Figure S2, the scintillating film is supported by a Si frame and by buffer layers of 20 nm of LaAlO3 and 5 unit cells of SrTiO3.37−39 A SEM image of four imaging “windows” fabricated on a silicon chip is shown in Figure 1b. As the electron beam scans above the encapsulated sample, light emitted from YAP:Ce on the Ce3+ allowed 4f−5d transitions is collected by a parabolic mirror (Figure 1c) and directed to a photomultiplier tube (PMT) or spectrometer. To generate an image, the signal from the PMT is correlated with the electron beam position. Due to the fast-scanning capability of the electron beam, images can be acquired at rates approaching those of other rapid imaging techniques, such as dark-field scattering microscopy or total internal reflection
beams with the noninvasiveness of optical approaches using very bright ultrathin scintillating films composed of ceriumdoped yttrium aluminum perovskite (YAlO3:Ce, or YAP:Ce).36 In our previous work, we prepared, characterized, and patterned YAP:Ce films on solid substrates, though at that point the presence of the underlying substrate precluded using these scintillating films for imaging. We found that the YAP:Ce films have a single CL emission peak centered at 370 nm, a 16 ns excitation lifetime, high spatial uniformity of CL, as we show in Supporting Information Figure S1, and a very low surface roughness. Here, we report the first realization of our noninvasive nanoimaging proposal by integrating this film into an imaging device and using this device to capture its first super-resolution imagesin this case, Al nanostructures and conjugated polymer films. Using the fast-scanning capability of the electron beam and the noninvasiveness that we achieve by exciting only the scintillator film and not the sample itself with the electrons, this CL-activated approach could be further extended for imaging biological processes in crowded environments with nanoscale resolution, high contrast, and video frame rates. B
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Figure 2. (a) Cathodoluminescence (CL)-activated image of Al nanostructures under the YAP:Ce scintillator film (scale bar, 1.0 μm). The inset shows a CL-activated image of a cluster of six Al nanostructures and the corresponding secondary electron (SE) image. (b) CL spectra of an area above an Al nanostructure (red) and an area with no Al nanostructure present (yellow), which correspond to the red and yellow boxes in part a, respectively. (c) CL-activated image of a single Al nanostructure. (d) Line-cut corresponding to the yellow line in the CL-activated image in panel c over the edge of a nanostructure, with a sigmoidal fit shown in red. The 80/20 width is 46 nm. (e) Photoluminescence (PL) lifetime measurements of YAP:Ce thin film (blue) and YAP:Ce thin film over an array of Al nanostructures (red). An additional 7.8 ns contribution to the PL decay emerges when the Al nanostructures are present in addition to the isolated YAP:Ce PL lifetime of 16.1 ns. The PL instrument response function (IRF) is indicated in gray.
fluorescence (TIRF) microscopy. Unlike these techniques, however, the electron-beam-induced lateral imaging resolution is not restricted by the diffraction limit and the resolution in the axial direction is defined by the near-field interactions between the film and the sample to provide far better background rejection than the evanescent field used in TIRF microscopy (70−250 nm). We call this imaging scheme “CL-activated imaging” because the low-energy electrons excite the Ce3+ dopants in the YAP:Ce film but do not penetrate into the sample beneath (Figure 1d). Instead, the optical excitations in the film noninvasively interact with the sample to produce contrast, which comes from differences in the photoemission rates of the Ce3+ dopants or the sample rather than differences in the number of scattered electrons. The differences in Ce3+ emission arise due to resonant energy transfer, which can in principle quench or enhance the Ce3+ luminescence, depending on the nature of the sample. Here, we first demonstrate in detail CL-activated imaging of Al nanostructures via Ce3+ luminescence enhancement and corroborate this contrast mechanism with simulations. Alternatively, when the Ce3+ luminescence is quenched via Förster resonance energy transfer (FRET) to a luminescent sample, the red-shifted emission from the sample can furthermore be correlated to the position of the electron beam to form its own image in tandem with the image formed from the scintillator luminescence. Second, we thus demonstrate unequivocally the noninvasiveness of CL-activated imaging on conjugated polymer films that are too delicate to be directly imaged with an electron beam. We do so by forming a CL-activated image of the polymer luminescence and by showing that its intensity variation is spatially anticorrelated to that of the tandem image formed from the scintillator CL. One component of the polymer film quenches Ce3+ luminescence
via FRET, as evidenced by its own spatially anticorrelated luminescence. First, we demonstrate CL-activated imaging of Al nanostructures encapsulated in SiO2 directly below the YAP:Ce film (Figure 1a). The Al nanostructures are 50 nm thick roughly triangular prisms and have a point-to-edge distance of approximately 270 nm, as determined before encapsulation by atomic force microscopy (AFM) (Figure 1e) and SEM (Figure 1f). They demonstrate the characteristic pattern of triangular nanostructures generated with nanosphere lithography. A CL-activated image resolving Al nanostructures embedded under the YAP:Ce with a 1.8 kV, 1.2 nA electron beam is shown in Figure 2a. The inset shows a higher magnification CL image of an arrangement of six nanostructures and the corresponding SEM secondary electron (SE) image. Because primary electrons do not reach the Al nanostructures, no additional detectable SEs are generated due to the presence of Al, and the Al nanostructures are not visible in the SE image. Spectra from the yellow- and red-boxed regions of the CL image are shown in Figure 2b. Clearly, when the electron beam is positioned above an Al nanostructure, the collected far-field radiation from the YAP:Ce film increases relative to an area with no Al. Our imaging approach, thus, enables nanoscale mapping of underlying samples by using the scintillating film as a high-resolution optical transducer that prevents electrons from reaching the sample. Characterizing this new imaging approach requires quantifying the resolution and contrast obtained in the CL-activated images. To determine our spatial resolution at 1.8 kV, we consider the YAP:Ce luminescence measured along the yellow line across the edge of an Al nanostructure (Figure 2c). The 80/20 width of the corresponding line-cut is 46 nm (Figure 2d), which is well below the diffraction limit. The luminescence intensity collected above the Al nanostructures is distinct from C
DOI: 10.1021/acs.nanolett.5b00716 Nano Lett. XXXX, XXX, XXX−XXX
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To mimic the fact that the electron beam activates a distribution of Ce3+ dopants, a weighted average of the radiative enhancement of a three-dimensional distribution of dipoles centered on a given pixel’s beam center position was calculated (see Supporting Information Figure S3). The resulting line-cut from the FDTD simulations matches both the width and shape of the experimental line-cut (Figure 3). The above experimental and theoretical observations are consistent with the following physical interpretation based on the fact that the detected luminescence from the Ce3+ dopants in the scanned excitation volume of the YAP:Ce film maps the electromagnetic density of states (LDOS) over top of the Al nanostructure.40−50 Depending on the location of a given Ce3+ relative to an Al nanostructure, its emission could be enhanced, quenched, or even possibly reflected. The total Ce 3+ luminescence collected over the course of a pixel dwell-time is altered in proximity to the Al nanostructures because a Ce3+ transition dipole induces a macroscopic polarization in the nearby metal nanostructure.40 The in-phase contribution to this polarization increases the Ce3+ emission rate and amplitude. A significant portion of the enhancement that we observe could arise from the edges and vertices of the Al nanostructure. For example, the shoulder peak on the right-hand side of the experimental line-cut in Figure 3 is reproduced in our FDTD simulations due to a higher degree of enhancement from the Al nanostructure vertex. Separately, a subpopulation of Ce3+ emitters closest to the Al surface may be quenched. Both enhancement and quenching effects could lead to the observed shortened lifetime component in Figure 2e. The excitation energy in the near-field of some Ce3+ dopants further away (∼10−15 nm from the surface) could, in principle, be radiated and detected in the far field through “reflection” off of the Al surface due to induced optical oscillations of bound electrons, though such reflection cannot entirely account for the observed increase in detected CL over top of the Al structures or for the shortened PL lifetime component. The enhancement along the perimeter of the nanostructure is well-resolved in the FDTD simulations of individual Ce3+ emitters nearby the nanostructure (Supporting Information Figure S3); the enhancement once averaged over the multiple emitters present in the electron beam excitation volume and plotted in Figure 3 is less pronounced along the perimeter of the structure. Yet, the fact that the simulations agree well with our experimental observations and also resolve the CL enhancement along the perimeter of the nanostructure shows that reflection alone cannot be responsible for the contrast in our experimental CLactivated images. In any case, each of these possible enhancement, quenching, and “reflective” contributions to the observed signal is a manifestation of a near-field interaction between the Al nanostructure sample and the scintillating film that can give rise to subdiffraction imaging contrast. The near-field imaging mechanism also explains the measured spatial resolution, which results from convolving the electron beam excitation volume in the scintillator (∼20 nm across36) with the distance-dependent profile of the electromagnetic interaction between the nanostructure and a single Ce3+ emitter, as shown in Supporting Information Figure S3. As such, the measurable resolution for imaging this particular type of nanostructure is likely lower than the resolution achievable with other samples. Regardless, our resolution, signal-to-noise ratio, and contrast, in addition to the advantage of avoiding mechanical scanning, demonstrate that CL-activation is a promising alternative to other imaging approaches.
that collected above SiO2, so that the nanostructures are easily discerned from background luminescence. The combination of high resolution and high contrast thus demonstrates the viability of CL-activated imaging. The high contrast and resolution of our imaging platform results from the near-field interactions between Ce3+ dopants in the scintillating film and the adjacent Al nanostructures. We elucidate the nature of these interactions with a combination of time-resolved photoluminescence (PL), spectrally resolved CL, control imaging measurements, and finite-difference timedomain (FDTD) simulations. As shown in Figure 2e, an additional 7.8 ns contribution to the PL decay of YAP:Ce (typically 16.1 ns) emerges when the Al nanostructures are present. When the electron beam is positioned over an Al nanostructure in our imaging configuration, the luminescence approximately doubles (Figure 2b). Furthermore, a “control” imaging chip identical to that shown in Figure 1a, but without YAP:Ce, was also studied. No detectable luminescence from the Al nanostructures was observed up to an accelerating voltage of 5.0 kV, implying that the observed enhancement at 1.8 kV cannot be caused by direct electron excitation. In addition to these experimental observations, a model of a CL line-cut across an entire Al nanostructure, obtained with FDTD simulations, agrees with the corresponding experimental line-cut through our CL image (Figure 3). The orientation of
Figure 3. Experimental line-cut from edge to tip of an Al nanostructure (green) and the simulated FDTD line cut (blue). The dots on the simulated line-cut indicate the location of the center of the modeled dipole distribution for each position on the line-cut. The yellow line in the inset CL-activated image indicates the position of the experimental line-cut. The directionality of the line-cut is known due to the relative position of the metal nanoparticles with respect to one another in nanosphere lithography, as shown in Figure 1.
the nanostructure in the CL image was determined based on its position with respect to adjacent Al nanostructures. To simulate the experimental line-cut, we modeled the luminescence of a single radiating Ce3+ dipole placed above an Al nanostructure, and we scanned its position in 20 nm increments along a 750 nm long line running through the edge, center, and tip of the nanostructure at a range of heights above the scintillator-sample interface. At each point, and for three orthogonal dipole orientations, the radiative enhancement factor was calculated based on the radiative power flux through a surface positioned above the dipole to mimic light collection. D
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Figure 4. (a) Plot of the enhancement factor versus the electron beam accelerating voltage. As the accelerating voltage increases, the enhancement factor increases linearly (1.0−2.0 kV) and then plateaus (2.0−3.0 kV). (b)−(e) Representative images at 1.0, 1.6, 2.2, and 2.8 kV, respectively. (f) Monte Carlo simulations of electron trajectories at 1.0 kV, 1.8 kV, and 2.2 kV. At 1.8 kV, the enhancement factor is within error of the plateau value and the electrons do not penetrate past the interface between the YAP:Ce film and the Al nanostructures.
Figure 5. (a) Schematic showing the geometry for imaging a luminescent polymer film. (b) CL-activated image of scintillator film quenching due to the presence of the adjacent luminescent polymer sample. (c) CL-activated image of emission from the luminescent polymer (PFO). The anticorrelation of panels b and c demonstrates that quenching of the scintillator film in the presence of the polymer is proportional to energy transfer from the scintillator film to the polymer, resulting in polymer emission. (d) YAP:Ce film CL emission is shown in purple, the overlapping absorption of the PFO luminescent polymer is shown in blue, and its photoluminescence (PL) emission is shown in green. (e) Fluorescence lifetime of YAP:Ce (purple) decreasing in the presence of PFO (green). The instrument response function (IRF) is shown in gray. (f) line-cut of the image in panel b at the position of the yellow line has an 80/20 width of 68 nm.
nanostructures were acquired at accelerating voltages ranging from 1.0 to 3.0 kV. A plot of the CL enhancement factor versus accelerating voltage is shown in Figure 4a, with representative images in Figure 4b−e at 1.0 kV, 1.6 kV, 2.2 kV, and 2.8 kV, respectively. Here, the enhancement factor represents the Ce3+ CL intensity measured above the Al nanostructures as compared to baseline YAP:Ce CL levels at each measured
Further optimization of our nanoimaging could be achieved by adjusting the electron accelerating voltage and scintillator film thickness, because both impact the contrast and resolution of the acquired images through the spatial distribution of Ce3+ excitations in the scintillator. To characterize how the imaging contrast depends on Ce3+ activation at different electron penetration depths into the YAP:Ce film, images of Al E
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variations observed on this length scale could be due to ripples in the film surface or due to the presence of F8BT, which could also act as a FRET acceptor to PFO. Importantly, these images also demonstrate that CL-activated imaging is nondamaging to soft materials. By contrast, attempts at direct imaging of similar polymer films with an electron beam do not yield CL images due to instantaneous bleaching of the organic chromophores upon interaction with the electrons. In sum, we have introduced subdiffraction optical imaging using a CL-activated near-field scanning optical microscopy that employs the nanoscale excitation volume generated by a lowenergy electron beam in a thin YAP:Ce scintillator film. We have demonstrated its capability by mapping the luminescence enhancement of Ce3+ dopants in the scintillator film due to the presence of Al nanostructures with 46 nm resolution and by mapping the Ce3+ luminescence quenching in tandem with the luminescence of conjugated polymers with 68 nm resolution. Because CL-activated nanoimaging maps how the presence of the sample affects the emission of nearby Ce3+ emitters, this approach provides information that is complementary to typical electron imaging techniques used to study the optical properties of nanostructures. For example, the more highly spatially resolved electron energy loss spectroscopy and direct CL measurements both rely on direct interactions between electrons and the sample. We have shown, however, that CLactivated imaging can additionally resolve nanoscale features in materials that cannot be imaged using these traditional electron microscopies due to damage induced by direct electron excitation. In particular, we have demonstrated that the Ce3+ luminescent centers excited by an electron beam can transfer energy via FRET to samples with endogenous chromophores such as polymer blends, whose red-shifted emission is correlated with the position of the electron beam to form an image. Furthermore, our CL-activated approach can be used for nanoimaging of a large class of samples with endogenous chromophores, such as soft, functional materials found, for example, in organic and hybrid organic−inorganic solar cells and light emitting diodes. This capability puts it at a great advantage over traditional super-resolution fluorescence microscopy, whose average spatial resolutions are comparable, in situations where one cannot use exogenous fluorescent or photoswitchable labels. Moreover, by fabricating free-standing membranes of YAP:Ce, we could create an imaging device that also serves as a liquid sample cell containing biomolecules, where the YAP:Ce free-standing film would act as a barrier between the vacuum environment of the SEM and the aqueous environment of the sample. In this fashion, the YAP:Ce film could FRET to the sample’s spectrally compatible fluorescent labels in the same way that we have shown with the polymer film. Specifically, small dye molecules or fluorescent proteins conjugated to the biomolecules of interest should couple to the scintillator film in precisely the same way as do the chromophores in the polymer film. Using this cell in combination with the fast-scanning capability of the electron beam, one could resolve the real-time dynamics of biomolecular interactions under physiological conditions and at their characteristic length scales. For example, the nanoscale dynamics of deoxyribonucleic acid (DNA) repair could be visualized by tethering DNA to the surface of the scintillator film and imaging a labeled protein diffusing in one-dimension along the DNA. One could also observe the two-dimensional diffusion of labeled membrane proteins on a crowded lipid
accelerating voltage (see Supporting Information Figure S4). The enhancement factor increases linearly as the accelerating voltage increases from 1.0 to 2.0 kV, and it plateaus beyond 2.0 kV. By consulting Monte Carlo simulations51 (Figure 4f) of elastic and inelastic electron scattering as a function of accelerating voltage, we attribute the linear increase in Figure 4a to a progressive increase in the number of activated Ce3+ emitters that are located close enough to the Al nanostructure to be enhanced. More of these dopants are reached and excited as the electrons penetrate deeper into the film. Thus, between 1.0 and 2.0 kV the signal-to-noise ratio and enhancement factor both improve. Yet, beyond 2.0 kV, the enhancement factor plateaus because there are no additional Ce3+ to be excited. The plateau could also be caused by a balance of CL enhancement of some Ce3+ emitters and CL quenching of others closer to the Al surface. In any case, the Monte Carlo simulation at 1.8 kV shows that very few electrons reach the Al even though they approach the interface. Thus, moving toward more delicate samples, 1.8 kV would be the ideal accelerating voltage at this film thickness because the near-field interaction is maximized with a very low probability of electrons interacting directly with the sample (see Supporting Information Figures S5 and S6). Although our imaging study of Al nanostructures enables us to determine the measurement parameters required for noninvasive imaging, we chose to demonstrate unequivocally that CL-activated imaging can be successfully used to measure more delicate samples that could not be imaged with traditional CL microscopy. To show that CL-activated nanoimaging can also be performed on soft materials that are easily damaged by direct excitation with an electron beam, a conjugated polymer blend of polyfluorene (PFO) and poly(9,9-dioctylfluorene-altbenzothiadiazole) (F8BT), as one might find in bulk heterojunction organic solar cells,52 was encapsulated for imaging using a scheme similar to that used with the Al nanostructures (Figure 5a). Here, the imaging contrast mechanism relies on FRET between the scintillator (donor) and the PFO in the polymer film (acceptor), which is facilitated by physical proximity and by strong spectral overlap between donor emission and acceptor absorption (Figure 5d). This near-field, nonradiative transfer of excitation energy from donor to acceptor requires that the amount of luminescence originating from a spot in the polymer film should be accompanied by a proportional decrease in the amount of luminescence from the adjacent spot in the scintillator, which is precisely what we observe. By recording in parallel the emission of YAP:Ce (Figure 5b) and the red-shifted emission of PFO (Figure 5c), we find that the spatial maps of polymer and scintillator emission are anticorrelated in intensity. Areas of low CL emission in Figure 5b appear where the scintillator film can transfer significant excitation energy to nearby PFO; the corresponding regions in Figure 5c PFO channel are very bright. In contrast, high intensity YAP:Ce luminescence areas show that PFO is too far from the YAP:Ce surface for significant amounts of energy transfer to occur, and correspond to regions of low intensity in Figure 5c. Our contrast mechanism interpretation is supported by the time-resolved PL measurements in Figure 5e that show that the YAP:Ce lifetime is reduced from 16.5 to 10.2 ns when PFO is applied to the surface. Additionally, a line-cut across a gradient in luminescence in Figure 5b has an 80/20 width of 68 nm (Figure 5f), which is well below the diffraction limit and is thus consistent with a near-field, rather than far-field, interaction between the scintillator and polymer film. The intensity F
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membrane or the light-induced reorganization of protein complexes in a densely packed thylakoid membrane to unravel the regulation of energy flow in photosynthesis.
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REFERENCES
(1) Betzig, E.; Patterson, G. H.; Sougrat, R.; Lindwasser, O. W.; Olenych, S.; Bonifacino, J. S.; Davidson, M. W.; Lippincott-Schwartz, J.; Hess, H. F. Science 2006, 313 (5793), 1642−1645. (2) Rust, M. J.; Bates, M.; Zhuang, X. Nat. Methods 2006, 3 (10), 793−796. (3) Hess, S. T.; Girirajan, T. P. K.; Mason, M. D. Biophys. J. 2006, 91 (11), 4258−4272. (4) Pavani, S. R. P.; Thompson, M. A.; Biteen, J. S.; Lord, S. J.; Liu, N.; Twieg, R. J.; Piestun, R.; Moerner, W. E. Proc. Natl. Acad. Sci. U.S.A. 2009, 106 (9), 2995−2999. (5) Hell, S. W. Science 2007, 316 (5828), 1153−1158. (6) Gustafsson, M. G. L. Proc. Natl. Acad. Sci. U.S.A. 2005, 102 (37), 13081−13086. (7) Garcia-Parajo, M. F. Nat. Photonics 2008, 2 (4), 201−203. (8) Kawata, S.; Inouye, Y.; Verma, P. Nat. Photonics 2009, 3 (7), 388−394. (9) Novotny, L.; van Hulst, N. Nat. Photonics 2011, 5 (2), 83−90. (10) Lewis, A.; Isaacson, M.; Harootunian, A.; Muray, A. Ultramicroscopy 1984, 13 (3), 227−231. (11) Pohl, D. W.; Denk, W.; Lanz, M. Appl. Phys. Lett. 1984, 44 (7), 651−653. (12) Betzig, E.; Trautman, J. K.; Harris, T. D.; Weiner, J. S.; Kostelak, R. L. Science 1991, 251 (5000), 1468−1470. (13) Zenhausern, F.; O’Boyle, M. P.; Wickramasinghe, H. K. Appl. Phys. Lett. 1994, 65 (13), 1623−1625. (14) Hartschuh, A.; Sánchez, E. J.; Xie, X. S.; Novotny, L. Phys. Rev. Lett. 2003, No. 9, 095503−1. (15) Kühn, S.; Håkanson, U.; Rogobete, L.; Sandoghdar, V. Phys. Rev. Lett. 2006, 97 (1), 017402. (16) Neacsu, C. C.; Berweger, S.; Olmon, R. L.; Saraf, L. V.; Ropers, C.; Raschke, M. B. Nano Lett. 2010, 10 (2), 592−596. (17) Weber-Bargioni, A.; Schwartzberg, A.; Cornaglia, M.; Ismach, A.; Urban, J. J.; Pang, Y.; Gordon, R.; Bokor, J.; Salmeron, M. B.; Ogletree, D. F.; Ashby, P.; Cabrini, S.; Schuck, P. J. Nano Lett. 2011, 11 (3), 1201−1207. (18) Zhu, Y.; Inada, H.; Nakamura, K.; Wall, J. Nat. Mater. 2009, 8 (10), 808−812. (19) Yacobi, B. G.; Holt, D. B. Cathodoluminescence microscopy of inorganic solids; Plenum Press: New York, 1990. (20) Yamamoto, N.; Araya, K.; García de Abajo, F. J. Phys. Rev. B 2001, 64 (20), 205419. (21) Vesseur, E. J. R.; de Waele, R.; Kuttge, M.; Polman, A. Nano Lett. 2007, 7 (9), 2843−2846. (22) Knight, M. W.; Liu, L.; Wang, Y.; Brown, L.; Mukherjee, S.; King, N. S.; Everitt, H. O.; Nordlander, P.; Halas, N. J. Nano Lett. 2012, 12 (11), 6000−6004. (23) Sapienza, R.; Coenen, T.; Renger, J.; Kuttge, M.; van Hulst, N. F.; Polman, A. Nat. Mater. 2012, 11 (9), 781−787. (24) Kociak, M.; Stéphan, O. Chem. Soc. Rev. 2014, 43 (11), 3865− 3883. (25) Coenen, T.; Arango, F. B.; Koenderink, A. F.; Polman, A. Nat. Commun. 2014, 5, 3250. (26) Haegel, N. M.; Chisholm, D. J.; Cole, R. A. J. Cryst. Growth 2012, 352 (1), 218−223. (27) Haegel, N. M. Nanophotonics 2013, 3 (1−2), 75−89. (28) Zagonel, L. F.; Mazzucco, S.; Tencé, M.; March, K.; Bernard, R.; Laslier, B.; Jacopin, G.; Tchernycheva, M.; Rigutti, L.; Julien, F. H.; Songmuang, R.; Kociak, M. Nano Lett. 2011, 11 (2), 568−573. (29) Hayes, T. L.; Pease, R. F. Adv. Biol. Med. Phys. 1967, 12, 85− 137. (30) Herbst, R.; Hoder, D. Scanning 1978, 1 (1), 35−41. (31) Niioka, H.; Furukawa, T.; Ichimiya, M.; Ashida, M.; Araki, T.; Hashimoto, M. Appl. Phys. Express 2011, 4 (11), 112402. (32) Glenn, D. R.; Zhang, H.; Kasthuri, N.; Schalek, R.; Lo, P. K.; Trifonov, A. S.; Park, H.; Lichtman, J. W.; Walsworth, R. L. Sci. Rep. 2012, 2, 865. (33) Furukawa, T.; Niioka, H.; Ichimiya, M.; Nagata, T.; Ashida, M.; Araki, T.; Hashimoto, M. Opt. Express 2013, 21 (22), 25655.
ASSOCIATED CONTENT
S Supporting Information *
The Supporting Information contains a quantitative measure of the brightness uniformity of the scintillator thin film, a signalto-noise ratio discussion, a description of the materials and methods, an explanation of the finite-difference time-domain (FDTD) calculations, a discussion regarding how we calculated our experimental CL enhancement factor, and a discussion of balancing the imaging brightness, damage, and resolution as a function of accelerating voltage. This material is available free of charge via the Internet at http://pubs.acs.org.
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AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected]. Present Address ○
KLA Tencor, 1 Technology Drive, Milpitas, California 95035, United States.
Author Contributions
N.S.G. conceived of and supervised the project. C.G.B., C.L.H., D.M.K., and N.S.G. designed the research. C.G.B. grew the YAP:Ce films, fabricated the Al nanostructures and the imaging device for the Al nanostructures and polymer blends, acquired and analyzed SE and CL images and PL lifetime measurements and performed CASINO and FDTD simulations. C.L.H. performed the PECVD nitride deposition step of the imaging device fabrication. Z.W. grew the Si/SrTiO3/LaAlO3 substrates. J.T.P. acquired AFM images of the Al nanostructures. C.G.B., C.L.H., and J.T.P. analyzed the data. D.G.S. advised on SrTiO3/ LaAlO3 film preparation. C.G.B. and N.S.G. wrote the manuscript, and all authors revised and approved the manuscript. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS YAP:Ce film deposition and CL characterization were supported by the National Science Foundation under Grant Number 1152656. Nanofabrication was supported by the Chemical Sciences, Geosciences and Biosciences Division, Office of Basic Energy Sciences, Office of Science, U.S. Department of Energy, FWP number SISGRN. Devices were fabricated both at the Marvell Nanofabrication Laboratory and Biomolecular Nanotechnology Center at UC Berkeley. CL and time-resolved fluorescence at the LBL Molecular Foundry were performed as part of the Molecular Foundry user program, supported by the Office of Science, Office of Basic Energy Sciences, of the U.S. Department of Energy under Contract No. DE-AC02-05CH11231. Z.W. and D.G.S. acknowledge support under the AFOSR Grant No. FA9550-10-1-0123, C.G.B. acknowledges an NSF Graduate Research Fellowship (DGE 1106400) and N.S.G. acknowledges a David and Lucile Packard Fellowship for Science and Engineering. We thank S. Aloni and D. F. Ogletree for providing technical and theoretical advice, and we thank R. Ramesh for access to PLD facilities. G
DOI: 10.1021/acs.nanolett.5b00716 Nano Lett. XXXX, XXX, XXX−XXX
Letter
Nano Letters (34) Yoon, H. P.; Lee, Y.; Bohn, C. D.; Ko, S.-H.; Gianfrancesco, A. G.; Steckel, J. S.; Coe-Sullivan, S.; Talin, A. A.; Zhitenev, N. B. AIP Adv. 2013, 3 (6), 062112. (35) Inami, W.; Nakajima, K.; Miyakawa, A.; Kawata, Y. Opt. Express 2010, 18 (12), 12897−12902. (36) Kaz, D. M.; Bischak, C. G.; Hetherington, C. L.; Howard, H. H.; Marti, X.; Clarkson, J. D.; Adamo, C.; Schlom, D. G.; Ramesh, R.; Aloni, S.; Ogletree, D. F.; Ginsberg, N. S. ACS Nano 2013, 7 (11), 10397−10404. (37) Kumaran, R.; Webster, S. E.; Penson, S.; Li, W.; Tiedje, T. J. Cryst. Growth 2009, 311 (7), 2191−2194. (38) Warusawithana, M. P.; Cen, C.; Sleasman, C. R.; Woicik, J. C.; Li, Y.; Kourkoutis, L. F.; Klug, J. A.; Li, H.; Ryan, P.; Wang, L.-P.; Bedzyk, M.; Muller, D. A.; Chen, L.-Q.; Levy, J.; Schlom, D. G. Science 2009, 324 (5925), 367−370. (39) Warusawithana, M. P.; Richter, C.; Mundy, J. A.; Roy, P.; Ludwig, J.; Paetel, S.; Heeg, T.; Pawlicki, A. A.; Kourkoutis, L. F.; Zheng, M.; Lee, M.; Mulcahy, B.; Zander, W.; Zhu, Y.; Schubert, J.; Eckstein, J. N.; Muller, D. A.; Hellberg, C. S.; Mannhart, J.; Schlom, D. G. Nat. Commun. 2013, 4, 2351. (40) Gersten, J.; Nitzan, A. J. Chem. Phys. 1981, 75 (3), 1139−1152. (41) Moskovits, M. Rev. Mod. Phys. 1985, 57 (3), 783−826. (42) Ebenstein, Y.; Mokari, T.; Banin, U. J. Phys. Chem. B 2004, 108 (1), 93−99. (43) Lakowicz, J. R. Anal. Biochem. 2005, 337 (2), 171−194. (44) Anger, P.; Bharadwaj, P.; Novotny, L. Phys. Rev. Lett. 2006, 96 (11), 113002. (45) Pompa, P. P.; Martiradonna, L.; Torre, A. D.; Sala, F. D.; Manna, L.; Vittorio, M. D.; Calabi, F.; Cingolani, R.; Rinaldi, R. Nat. Nanotechnol. 2006, 1 (2), 126−130. (46) Willets, K. A.; Van Duyne, R. P. Annu. Rev. Phys. Chem. 2007, 58 (1), 267−297. (47) Kinkhabwala, A.; Yu, Z.; Fan, S.; Avlasevich, Y.; Müllen, K.; Moerner, W. E. Nat. Photonics 2009, 3 (11), 654−657. (48) Masiello, D. J.; Schatz, G. C. J. Chem. Phys. 2010, 132 (6), 064102. (49) Munechika, K.; Chen, Y.; Tillack, A. F.; Kulkarni, A. P.; Plante, I. J.-L.; Munro, A. M.; Ginger, D. S. Nano Lett. 2010, 10 (7), 2598− 2603. (50) Dionne, J. A.; Atwater, H. A. MRS Bull. 2012, 37 (08), 717− 724. (51) Drouin, D.; Couture, A. R.; Joly, D.; Tastet, X.; Aimez, V.; Gauvin, R. Scanning 2007, 29 (3), 92−101. (52) Yu, G.; Gao, J.; Hummelen, J. C.; Wudl, F.; Heeger, A. J. Science 1995, 270 (5243), 1789−1791.
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