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Optics Express

Optics Express

  • Editor: Andrew M. Weiner
  • Vol. 21, Iss. 23 — Nov. 18, 2013
  • pp: 28198–28218
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Diffraction-unlimited optical imaging of unstained living cells in liquid by electron beam scanning of luminescent environmental cells

Hideki T. Miyazaki, Takeshi Kasaya, Taro Takemura, Nobutaka Hanagata, Takeshi Yasuda, and Hiroshi Miyazaki  »View Author Affiliations


Optics Express, Vol. 21, Issue 23, pp. 28198-28218 (2013)
http://dx.doi.org/10.1364/OE.21.028198


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Abstract

An environmental cell with a 50-nm-thick cathodolumi-nescent window was attached to a scanning electron microscope, and diffraction-unlimited near-field optical imaging of unstained living human lung epithelial cells in liquid was demonstrated. Electrons with energies as low as 0.8 – 1.2 kV are sufficiently blocked by the window without damaging the specimens, and form a sub-wavelength-sized illumination light source. A super-resolved optical image of the specimen adhered to the opposite window surface was acquired by a photomultiplier tube placed below. The cells after the observation were proved to stay alive. The image was formed by enhanced dipole radiation or energy transfer, and features as small as 62 nm were resolved.

© 2013 Optical Society of America

1. Introduction

Optical microscopy, which provides informative images of various objects in diverse environments, has played a vital role in many fields, particularly biology [1

1. S. Inoué and K. R. Spring, Video Microscopy, the Fundamentals, 2 (Plenum, 1997). [CrossRef]

]. Phase contrast microscopy is indispensable for observing living cells in liquid without staining. Fluorescence microscopy, which visualizes specific molecules or organelles, has also yielded an enormous number of biological results. However, the imaging resolution of all these techniques is limited by diffraction. Electron microscopy provides a resolution [2

2. L. Reimer, Transmission Electron Microscopy, 4 (Springer, 1997). [CrossRef]

, 3

3. L. Reimer, Scanning Electron Microscopy, 2 (Springer, 1998). [CrossRef]

] far beyond the diffraction limit of optical microscopy, but the vacuum environment surrounding the specimen precludes observation of wet, living cells. Furthermore, complicated sample preparation is necessary in order to maintain the natural state of the objects, and important signals such as fluorescence, light absorption, and refractive index are lost.

To compensate for the drawbacks of each type of microscopy, labeling techniques and systems have been improved to allow correlated observations of a single specimen by optical and electron microscopy [4

4. J. E. Mazurkiewicz and P. K. Nakane, “Light and electron microscopic localization of antigens in tissues embedded in polyethylene glycol with a peroxidase-labeled antibody method,” J. Histochem. Cytochem. 20, 969–974 (1972). [CrossRef] [PubMed]

]. New technologies have also been intensively developed in each field.

In optical microscopy, the near-field scanning optical microscope (NSOM) broke the diffraction limit [5

5. D. W. Pohl, W. Denk, and M. Lanz, “Optical stethoscopy: image recording with resolution λ/20,” Appl. Phys. Lett. 44, 651–653 (1984). [CrossRef]

,6

6. A. Lewis, M. Isaacson, A. Harootunian, and A. Muray, “Development of a 500 Å spatial resolution light microscope,” Ultramicroscopy 13, 227–232 (1984). [CrossRef]

], and has become an established tool for visualizing the localized light in nanostructures [7

7. T. Saiki, K. Nishi, and M. Ohtsu, “Low temperature near-field photoluminescence spectroscopy of InGaAs single quantum dots,”Jpn. J. Appl. Phys. 37, 1638–1642 (1998). [CrossRef]

]. However, the use of NSOMs to cells in liquid is limited [8

8. M. Koopman, A. Cambi, B. I. de Bakker, B. Joosten, C. G. Figdor, N. F. van Hulst, and M. F. Garcia-Parajo, “Near-field scanning optical microscopy in liquid for high resolution single molecule detection on dendritic cells,” FEBS Lett. 573, 6–10 (2004). [CrossRef] [PubMed]

,9

9. C. Höppener and L. Novotny, “Imaging of membrane proteins using antenna-based optical microscopy,” Nanotechnology 19, 384012 (2008). [CrossRef] [PubMed]

] due to the difficulty in controlling the distance of the probe in a liquid and observations are usually based on fluorescent labeling. Tip-enhanced Raman scattering [10

10. N. Hayazawa, Y. Inouye, Z. Sekkat, and S. Kawata, “Metallized tip amplification of near-field Raman scattering,” Opt. Commun. 183, 333–336 (2000). [CrossRef]

] does not require staining; however, its application in biology is still in its preliminary stages [11

11. R. Böhme, M. Richter, D. Cialla, P. Rösch, V. Deckert, and J. Popp, “Towards a specific characterisation of components on a cell surface – combined TERS – investigations of lipids and human cells,” J. Raman Spectrosc. 40, 1452–1457 (2009). [CrossRef]

].

In this decade, various optical far-field nanoscopy techniques based on nonlinearity in fluorescence [12

12. T. A. Klar, S. Jakobs, M. Dyba, A. Egner, and S. W. Hell, “Fluorescence microscopy with diffraction resolution barrier broken by stimulated emission,” Proc. Natl. Acad. Sci. U. S. A. 97, 8206–8210 (2000). [CrossRef] [PubMed]

, 13

13. M. G. L. Gustafsson, “Nonlinear structured-illumination microscopy: wide-field fluorescence imaging with theoretically unlimited resolution,” Proc. Natl. Acad. Sci. U. S. A. 102, 13081–13086 (2005). [CrossRef] [PubMed]

] or single-molecule detection [14

14. E. Betzig, G. H. Patterson, R. Sougrat, O. W. Lindwasser, S. Olenych, J. S. Bonifacino, M. W. Davidson, J. Lippincott-Schwartz, and H. F. Hess, “Imaging intracellular fluorescent protains at nanometer resolution,” Science 313, 1642–1645 (2006). [CrossRef] [PubMed]

, 15

15. M. J. Rust, M. Bates, and X. Zhuang, “Sub-diffraction-limit imaging by stochastic optical reconstruction microscopy (STORM),” Nat. Methods 3, 793–795 (2006). [CrossRef] [PubMed]

] have been developed, and three-dimensional diffraction-unlimited observation of living cells in liquid through the use of traditional optical systems based on lenses has been realized [16

16. S. W. Hell, “Far-field optical nanoscopy,” Science 316, 1153–1158 (2007). [CrossRef] [PubMed]

]. However, these techniques, in principle, require fluorescent labeling and the development of far-field nanoscopy of unstained living cells is currently among the most active areas of research [17

17. Y. Cotte, F. Toy, P. Jourdain, N. Pavillon, D. Boss, P. Magistretti, P. Marquet, and C. Depeursinge, “Marker-free phase nanoscopy,” Nat. Photonics 7, 113–117 (2013). [CrossRef]

, 18

18. P. Wang, M. N. Slipchenko, J. Mitchell, C. Yang, E. O. Potma, X. Xu, and J. X. Cheng, “Far-field imaging of non-fluorescent species with subdiffraction resolution,” Nat. Photonics 7, 449–453 (2013). [CrossRef]

].

Another outstanding achievement in electron microscopy is the development of environmental cells (ECs). The observation of wet, unstained, and unfixed biological objects has long been an important issue [27

27. D. F. Parsons, V. R. Matricardi, R. C. Moretz, and J. N. Turner, “Electron microscopy and diffraction of wet unstained and unfixed biological objects,” Adv. Biol. Med. Phys. 15, 161–270 (1974). [PubMed]

], but the progress in this past decade has been remarkable. ECs have one or two thin membranes that separate the atmosphere around the specimen from the vacuum, and successful observations of objects in liquids by detecting transmitted electrons, scattered electrons, or CL with transmission electron microscopes (TEM) [28

28. K.-L. Liu, C.-C. Wu, Y.-J. Huang, H.-L. Peng, W.-Y. Chang, P. Chang, L. Hsu, and T.-R. Yew, “Novel microchip for in situ TEM imaging of living organisms and bio-reactions in aqueous conditions,” Lab Chip 8, 1915–1921 (2008). [CrossRef] [PubMed]

], scanning transmission electron microscopes (STEM) [29

29. N. de Jonge, D. B. Peckys, G. J. Kremers, and D. W. Piston, “Electron microscopy of whole cells in liquid with nanometer resolution,” Proc. Natl. Acad. Sci. U. S. A. 106, 2159–2164 (2009). [CrossRef] [PubMed]

], and scanning electron microscopes (SEM) [30

30. S. Thiberge, A. Nechushtan, D. Sprinzak, O. Gileadi, V. Behar, O. Zik, Y. Chowers, S. Michaeli, J. Schlessinger, and E. Moses, “Scanning electron microscopy of cells and tissues under fully hydrated conditions,” Proc. Natl. Acad. Sci. U. S. A. 101, 3346–3351 (2004). [CrossRef] [PubMed]

] have been reported [31

31. N. de Jonge and F. M. Ross, “Electron miocroscopy of specimens in liquid,” Nat. Nanotechnol. 6, 695–704 (2011). [CrossRef] [PubMed]

]. Even an inverted SEM, which is tailored for specimens in liquid and does not require an enclosed EC, has been developed [32

32. H. Nishiyama, M. Suga, T. Ogura, Y. Maruyama, M. Koizumi, K. Mio, S. Kitamura, and C. Sato, “Atmospheric scanning electron microscope observes cells and tissues in open medium through silicon nitride film,” J. Struct. Biol. 169, 438–449 (2010). [CrossRef] [PubMed]

]. Damage to the object imposes serious limitations on the electron microscopy of biological specimens in liquid [33

33. V. E. Cosslett, “Radiation damage in the high resolution electron microscopy of biological materials: a review,” J. Microsc. 113, 113–129 (1978). [CrossRef] [PubMed]

]. Because the image is based on electron absorption, scattering, or CL by the specimen, exposure of the specimen to the electrons is inevitable. While it is demonstrated that bacteria remained viable after TEM observation with a minimized electron dose [28

28. K.-L. Liu, C.-C. Wu, Y.-J. Huang, H.-L. Peng, W.-Y. Chang, P. Chang, L. Hsu, and T.-R. Yew, “Novel microchip for in situ TEM imaging of living organisms and bio-reactions in aqueous conditions,” Lab Chip 8, 1915–1921 (2008). [CrossRef] [PubMed]

], it is also reported that the acquisition of a single STEM image is enough to kill cells even if the dose is kept below conventionally known threshold [34

34. D. B. Peckys, P. Mazur, K. L. Gould, and N. de Jonge, “Fully hydrated yeast cells imaged with electron microscopy,” Biophys. J. 100, 2522–2529 (2011). [CrossRef] [PubMed]

].

Aiming at super-resolved optical imaging of living cells in liquid, techniques in optical microscopy and electron microscopy are being combined. Inami et al. proposed optical imaging of sub-diffraction-limit objects deposited on a luminescent membrane with an SEM [35

35. W. Inami, K. Nakajima, A. Miyakawa, and Y. Kawata, “Electron beam excitation assisted optical microscope with ultra-high resolution,” Opt. Express 18, 12897–12902 (2010). [CrossRef] [PubMed]

], and demonstrated the observation of biological cells in culture medium [36

36. Y. Nawa, W. Inami, A. Chiba, A. Ono, A. Miyakawa, Y. Kawata, S. Lin, and S. Terakawa, “Dynamic and high-resolution live cell imaging by direct electron beam excitation,” Opt. Express 20, 5629–5635 (2012). [CrossRef] [PubMed]

]. However, their approach requires a tailored inverted SEM and has not been applied to standard SEMs. Moreover, most of the electrons penetrate the membrane and impinge on the specimen. The images of the cells are based on CL directly excited by an EB as had been demonstrated by Thiberge et al. [30

30. S. Thiberge, A. Nechushtan, D. Sprinzak, O. Gileadi, V. Behar, O. Zik, Y. Chowers, S. Michaeli, J. Schlessinger, and E. Moses, “Scanning electron microscopy of cells and tissues under fully hydrated conditions,” Proc. Natl. Acad. Sci. U. S. A. 101, 3346–3351 (2004). [CrossRef] [PubMed]

], and it is not shown whether the cells stayed alive after the image acquisition. An X-ray microscope using characteristic X-rays from the entrance window of an environmental cell has also been developed [37

37. T. Ogura, “Measurement of the unstained biological sample by a novel scanning electron generation X-ray microscope based on SEM,” Biochem. Biophys. Res. Commun. 385, 624–629 (2009). [CrossRef] [PubMed]

], although it visualizes different information from optical microscopes and observation of specimen fully immersed in a liquid has not been shown.

In this paper, an environmental cell with a 50-nm-thick cathodoluminescent window is attached to a scanning electron microscope, and diffraction-unlimited near-field optical imaging of unstained living human lung epithelial cells in liquid is demonstrated. In addition to the cytoplasmic fibers and granules, toxic nanoparticles (NPs) taken up into cells are also observed. Electrons with energies as low as 0.8 – 1.2 kV are sufficiently blocked by the window without damaging the specimens, and form a sub-wavelength-sized illumination light source. A super-resolved optical image of the specimen adhered to the opposite window surface is acquired by a photomultiplier tube (PMT) placed below. The cells after the observation were proved to stay alive. The image is formed by enhanced dipole radiation or energy transfer, and features as small as 62 nm were resolved.

The remainder of this paper is organized as follows. In Sec. 2, the setup and principles of our microscope, which we call scanning electron optical microscope (SEOM), are introduced from the viewpoints of electron penetration and modification of dipole radiation. In Sec. 3, observations of NPs are described and actual image resolution is evaluated. In Sec. 4, the significance of observing unstained cells in cytotoxicity studies is described and images of unstained living cells, NPs taken up into the cells, and other representative results are presented. The limitations of the current SEOM system and its future outlook are discussed in Sec. 5, and the results are summarized in Sec. 6. Detailed information on materials and methods, and some supporting data are given in the Appendices.

2. System setup and principles

2.1. System setup

A schematic illustration and photographs of the constructed detection unit are shown in Figs. 1(a) and 1(b), respectively. This unit is attached to the specimen stage of a cold field-emission SEM (S-4200, Hitachi). The main body of the detection unit is a photon-counting PMT module (H10723-01MOD, Hamamatsu Photonics) equipped with a high-voltage power supply inside. The EC is positioned just above the photocathode and can be exchanged quickly through the airlock. The structure of the EC is shown in Fig. 1(c). The entrance window is a SiN membrane (details will be given later); the EB is incident on this window. The exit window, through which the light from the emitting layer deposited on the SiN membrane passes to the PMT, is a 0.25-mm-thick quartz disk. Both windows are hermetically glued to stainless supports and a small area between the windows is sealed by packing. These parts are joined together with two screws and can also be disassembled. In several experiments to confirm the device’s fundamental performance, the exit window was not used and the specimen was placed under vacuum.

Fig. 1 (a) Setup of SEOM. The EC placed under vacuum maintains an arbitrary environment between the entrance and exit windows. The cathodoluminescent entrance window is scanned with an EB, and light modulated by the specimen passes through the exit window and is detected by the PMT beneath. The SiN membrane is supported by a rigid Si frame. (b) Photographs of the detection unit attached to the specimen stage of the SEM. In the inset, the EC unit is removed and the photocathode can be seen. (c) Left: the entrance and the exit windows. Right: the exit side of the assembled state. There are two vent holes on either side of the exit window to drain excess liquid during assembly. After tightening the screws, each hole is sealed with polyimide film and an instant glue. (d) For culturing cells on the entrance window, a silicone ring is attached and filled with culture medium. Scale bars in (b) and (c): 10 mm.

The key component of the SEOM system is the entrance window of the EC. It should satisfy the following nine requirements. It must (1) be pinhole-free to retain the liquid or gas inside, (2) be mechanically strong to withstand a pressure difference of 1 atm, (3) convert electrons to photons efficiently, (4) be sufficiently thin to realize a small light source, (5) produce nanometrically uniform light emission, (6) possess a small exciton diffusion length, (7) have sufficient stopping power for the electrons, (8) be free from electric charging, and (9) provide sufficient adhesion for the cells.

After extensive trials, a SiN membrane (SiMPore) coated with poly(9-vinylcarbazole) (PVK) doped with 1 wt% coumarin 6 (C6) was employed as the entrance window. The EB is incident on the SiN side and the specimens are deposited on the PVK side. Two types of SiN membranes with different values of thickness tm were used: tm = 20 nm (0.5-mm square) and 15 nm (0.25-mm square). The typical values of thickness te for the PVK emitting film on the two types of SiN membrane are te = 30 nm and 35 nm, respectively; thus, in both cases, the total thickness tm + te is typically 50 nm. SiN membranes are pinhole-free and those with the above dimensions can withstand a pressure difference of 1 atm. C6-doped PVK (PVK:C6) is an efficient light emitter. The PVK matrix itself is a wide-bandgap blue-emitting polymer, but PVK:C6 exhibits green emission around 520 nm through Förster energy transfer to C6 [38

38. J. Kido, H. Shionoya, and K. Nagai, “Single-layer white light-emitting organic electroluminescent devices based on dye-dispersed poly(N-vinylcarbazole),” Appl. Phys. Lett. 67, 2281–2283 (1995). [CrossRef]

]. Thus, conversion of high-energy electrons to visible photons is expected. The exciton diffusion length of organic molecules is generally as short as 10 nm, and thus the emission region is dominated by electron diffusion [39

39. R. R. Lunt, N. C. Giebink, A. A. Belak, J. B. Benziger, and S. R. Forrest, “Exciton diffusion lengths of organic semiconductor thin films measured by spectrally resolved photoluminescence quenching,” J. Appl. Phys. 105, 053711 (2009). [CrossRef]

]. The EB can be stopped within the entrance window by choosing a sufficiently low acceleration energy E. Since the electrons diffuse nearly spherically [40

40. K. Kanaya and S. Okayama, “Penetration and energy-loss theory of electrons in solid targets,” J. Phys. D Appl. Phys. 5, 43–58 (1972). [CrossRef]

], the window thickness governs the size of the emission region. Therefore, the total thickness of the entrance windows tm + te was limited to 50 nm, and E was adjusted so that the electrons stopped within this thickness. Although SiN is easily charged in the low E region discussed in this work, the charging was found to be suppressed when the EC was filled with a liquid. NPs or biological cells can be deposited on the PVK surface by poly-l-lysine (PLL) coating. Thus, we obtained entrance windows satisfying all nine of the requirements.

The choice of an organic emitting layer should be regarded as tentative, because it is easily bleached by the EB and only a single image can be obtained from a particular area. We have tried various inorganic films to find an EB-durable emitting layer; however, emission was not laterally uniform and so they could not be used for our purposes (see Appendix B). In contrast, the uniformity of the emission from organic materials is excellent.

2.2. Electron penetration

To determine the optimum value of E, the electron transmission for entrance windows with different emitter thicknesses te was measured as a function of the electron energy E. The results are shown in Fig. 2(a). The transmittance is sensitive to E in the low energy region, and quickly increases after passing a threshold value, which strongly depends on the window thickness. The results obtained are very similar to those for carbon membranes [3

3. L. Reimer, Scanning Electron Microscopy, 2 (Springer, 1998). [CrossRef]

]. Since the observed current did not drop to zero, presumably due to some leakage current, the E value just before the sharp increase was regarded as the experimental threshold. The results of a Monte Carlo simulation [41

41. D. Drouin, A. R. Couture, D. Joly, X. Tastet, V. Aimez, and R. Gauvin, “CASINO V2.42—a fast and easy-to-use modeling tool for scanning electron microscopy and microanalysis users,” Scanning 29, 92–101 (2007). [CrossRef] [PubMed]

] are also shown in the inset; however, the agreement with the experiment is not remarkable. The minimum energy that shows a non-zero value was regarded as the theoretical threshold. These thresholds are slightly higher, and the increases in transmission are steeper than the experimental results. This discrepancy could be because low energy electrons (< 50 eV) are not included in the simulation. In Fig. 2(b), simulated distributions of the absorbed energy at the theoretical threshold for E are shown. In each case the distribution is nearly spherical and this determines the emission region. The electrons are stopped just inside the entrance window at the thresholds. Figure 2(c) shows the E dependence of the emission spectra from the entrance windows. The results are consistent with the measured transmittance. At sufficiently low values of E (0.6 kV), at which the electrons stay in the SiN membrane and do not reach the PVK film, a weak, broad emission from SiN is observed. When the electrons reach the PVK film (E ≥ 0.7 kV), strong emission from C6 centered at 520 nm is produced. In the observations described below, the acceleration energy E was set at the threshold, unless otherwise noted. Since tm and te differ for each specimen, the value of E also differs for each image.

Fig. 2 (a) Electron penetration through the entrance windows with tm = 20 nm and different te as a function of electron energy E. Experimental threshold energies are indicated by arrows. The threshold for tm = 15 nm and te = 35 nm was 0.8 kV (data not shown). Inset: magnification around the thresholds. The results of Monte Carlo simulation are shown as broken lines whose colors correspond to those of the experimental results. The theoretical thresholds are denoted by asterisks. (b) Simulated distributions of the absorbed energy at the theoretical threshold energies for te = 0, 30, and 60 nm (E = 0.9, 1.3, and 1.7 kV, respectively). These cases correspond to the experimental results for E = 0.6, 0.9, and 1.0 kV, respectively. (c) E dependence of the emission spectra of the entrance windows for tm = 20 nm and te = 30 nm. The spectra shown were acquired without the exit window, but those for enclosed cells filled with water were similar (data not shown).

2.3. Image formation

To clarify the principles of image formation and the limitations of the proposed method, expected images were simulated on the basis of the Maxwell equations. The specimen is represented as a dielectric sphere in water, and the change in the radiation from a dipole caused by the presence of the sphere in its near field was calculated by using the vector spherical wave expansion. Figure 3(a) shows the angular distribution of the radiation from a dipole located 10 nm above spheres with different refractive indices and sizes. Roughly speaking, the presence of the sphere enhances the radiation in the high angle region (θ ≃ 90°) [42

42. H. Mertens, A. F. Koenderink, and A. Polman, “Plasmon-enhanced luminescence near noble-metal nanospheres: comparison of exact theory and an improved Gersten and Nitzan model,” Phys. Rev. B 76, 115123 (2007). [CrossRef]

]. Therefore, the collection of high-angle emissions is important to obtain a high-contrast image. Geometric optics simulation (see Appendix C) shows that reflection off the sloped Si surfaces formed by the anisotropic etching surrounding the SiN membrane contributes to light collection in our setup. By moving the dipole horizontally, an isolated sphere is observed as a bright spot with a full width at half-maximum (FWHM) close to the diameter of the sphere, as shown in Fig. 3(b), left; for example, a 50-nm-diameter sphere is imaged with a FWHM of 85 nm. In this calculation, the spherical distribution of the electrons in the emitting layer is taken into account for accurate estimation of the real image. Since the diffraction limit 0.61 λ/NA (λ: wavelength, NA: numerical aperture) at λ = 550 nm is 240 nm (NA = 1.4 is assumed), this implies the possibility of diffraction-unlimited imaging. The image contrast, defined as the ratio of the intensity increase on the top of the sphere to the intensity far from the sphere, depends on the refractive index contrast (n2n1)/n1 and the diameter of the sphere. Figure 3(b) also shows the intensity profiles for two spheres in contact. It may be difficult to resolve two spheres with a diameter 2a = 100 nm, but spheres with 2a = 150 nm would be distinguished from each other. Figure 3(c) shows the change in the contrast according to the distance d from the emitting layer. The image contrast drops exponentially as the object gets further from the emitting surface with a decay distance of dλ/10, that is, ≃ 50 nm for visible light. Therefore, SEOM can visualize only structures near the adhesion interface of a specimen. This is very similar to conventional NSOM and total internal reflection fluorescence microscopy [43

43. T. Funatsu, Y. Harada, M. Tokunaga, K. Saito, and T. Yanagida, “Imaging of single fluorescent molecules and individual ATP turnovers by single myosin molecules in aqueous solution,” Nature 374, 555–559 (1995). [CrossRef] [PubMed]

]. When the medium is vacuum or air (n1 = 1), the image contrast increases, since it is proportional to the index contrast; however, the resolution remains similar to that in Fig. 3(b) (see Appendix D).

Fig. 3 (a) Angular distribution of the radiation at λ = 550 nm from a single dipole placed at a position Δz = 10 nm from the top of a sphere. As shown above the panel, the sphere is embedded in a medium with a refractive index n1. Upper: radiation modulated by spheres with a diameter 2a = 100 nm and various refractive indices n2 placed in water. We considered polystyrene (PS), SiO2, ZnO, and CuO. Lower: radiation modulated by PS spheres with various diameters placed in water. Even when the sphere is absent, a constant intensity is detected since a dipole always radiates homogeneously. The presence of the sphere enhances the radiation and gives a positive contrast. (b) Intensity profile at λ = 550 nm for the EB scanning. Left: single PS spheres. Right: two PS spheres in contact with various diameters. In the calculation hereafter, the distribution of the electrons is taken into account (red lower hemisphere above the panels). (c) Contrast change as a function of the distance d of the PS sphere from the surface of the emitting layer for several diameters and wavelengths. The dominant exponential terms are indicated by the lines. See Appendix A for details on the calculation.

3. Observation of NPs

To evaluate the imaging performance of the constructed SEOM system, NPs with various refractive indices discussed in Fig. 3 (PS, SiO2, ZnO, and CuO) with known shapes and sizes were observed. PS and SiO2 particles are uniform spheres, and are thus suitable for a quantitative discussion of the resolution. ZnO and CuO are important from the viewpoint of cytotoxicity, as discussed later. Since the resolution is not sensitive to the surrounding media, the evaluation was carried out for NPs under a vacuum. Figure 4 shows the images acquired by SEOM (called CL images hereafter), and the secondary electron (SE) images of the same area in the flipped entrance window observed afterward in conventional SEM mode. Representative intensity profiles are also shown and compared with those in the SE images.

Fig. 4 Images of NPs for evaluating the resolution. (a) – (d): CL (cathodoluminescence) images by SEOM system. (e) – (h): SE (secondary electron) images. (i) – (l): intensity profiles (red: CL, blue: SE) between the arrowheads in the CL/SE images for four particles. (a), (e), and (i): PS (diameter: 100 nm), (b), (f), and (j): SiO2 (100 nm), (c), (g), and (k): ZnO (< 100 nm), and (d), (h), and (l): CuO (< 50 nm). In the profiles, FWHM or peak distance is shown. In the SE images in (e) – (h), particles on the flipped entrance window were directly observed at E = 3 or 5 kV, as shown to the right of the images, and the flipped images are shown. Observation conditions of the CL images (E, pixel size Xp, dwell time Td) for each panel are (a) 0.9 kV, 20 nm, 10 ms, (b) 0.8 kV, 20 nm, 10 ms, (c) 0.8 kV, 20 nm, 10 ms, and (d) 1.0 kV, 15 nm, 5 ms. The CL images are lowpass filtered at a cutoff of 1/3 pixel−1. Though ZnO is a typical phosphor material, the CL from the ZnO NPs does not contribute to the image, since the EB is sufficiently blocked. Scale bars: 1 μm.

The uniform PS and SiO2 spheres form monolayer arrays and show fairly good correspondence between the CL and SE images. As shown in Figs. 4(a) and 4(i), an isolated PS sphere with a diameter 2a = 86 nm was observed as a bright spot with a FWHM of 108 nm, well below the diffraction limit. However, individual particles in clusters could not be resolved.

For SiO2 particles, clusters made of a few particles were visible, as shown in Fig. 4(b), but isolated spheres were hardly discernible. As has been predicted [Fig. 3(a)], the contrast for low-index SiO2 is small and the signal to noise (S/N) ratio is poor.

On the other hand, the ZnO particles in Fig. 4(c) are sharply imaged with high contrast, and features as small as 88 nm were visualized. However, the correspondence between the CL and SE images is not necessarily excellent. Some of the NPs [circles in Fig. 4(g)] do not appear in the CL image. ZnO particles are needle-shaped and the clusters are three-dimensionally stacked. By inspecting the sample from various directions by SEM, it was confirmed that the unobserved particles lie away from the emitting layer surface. This is consistent with the distance dependence shown in Fig. 3(c).

The ranks of the image contrasts are ZnO > PS > SiO2, which correspond to the relationships of the refractive indices. Quantitatively, the experimental contrasts in Figs. 4(a)–4(c) were higher than the theoretical predictions (data not shown). This discrepancy could be attributed to the oversimplified calculation model, and provides encouragement that observations of cells in water with a very small index contrast could be made.

The images for CuO, however, had unexpected properties. Although a particularly bright image for CuO was predicted [Fig. 3(a)], in the experimental image the NPs appeared as dark points on a lighter background (i.e., the contrast was opposite that for the other NPs) [Fig. 4(d)]. The bandgap of CuO, 1.56 eV, is smaller than that of C6, 2.3 eV [38

38. J. Kido, H. Shionoya, and K. Nagai, “Single-layer white light-emitting organic electroluminescent devices based on dye-dispersed poly(N-vinylcarbazole),” Appl. Phys. Lett. 67, 2281–2283 (1995). [CrossRef]

,44

44. K. Nakaoka, J. Ueyama, and K. Ogura, “Photoelectrochemical behavior of electrodeposited CuO and Cu2O thin films on conducting substrates,” J. Electrochem. Soc. 151, C661–C665 (2004). [CrossRef]

] so the negative contrast could be attributed to quenching by energy transfer from C6 to CuO. The images for CuO were very vivid, and features as narrow as 62 nm can be seen in Fig. 4(d). Thus, the contrast in an SEOM image of non-absorbing materials is formed by the difference between the refractive indices of the specimens and the surrounding medium, but for absorbing materials, the image contrast is based on the energy transfer.

4. Observation of unstained cells in liquid

As the industrial use of nanomaterials has increased, the safety of NPs, in particular of those taken up into the lung, has become a major concern. The toxicity of ZnO and CuO to human lung epithelial A549 cells has been revealed by a statistical viability assay of numbers of cells [45

45. M. Xu, D. Fujita, S. Kajiwara, T. Minowa, X. Li, T. Takemura, H. Iwai, and N. Hanagata, “Contribution of physicochemical characteristics of nano-oxides to cytotoxicity,” Biomaterials 31, 8022–8031 (2010). [CrossRef] [PubMed]

]. However, the uptake process and the subsequent aggregation of NPs in the cells are not fully understood. The difficulty lies in the fact that fluorescent labeling cannot be applied to the study of cytotoxicity. This is because the label attached to the NPs could itself change the interaction between the NPs and the cells. Although CuO NPs taken up into cells have successfully been observed by TEM in fixed, dehydrated, and ultrathin-sectioned specimens [46

46. N. Hanagata, F. Zhuang, S. Connolly, J. Li, N. Ogawa, and M. Xu, “Molecular responses of human lung epithelial cells to the toxicity of copper oxide nanoparticles inferred from whole genome expression analysis,”ACS Nano 5, 9326–9338 (2011). [CrossRef] [PubMed]

], more efficient techniques are required. Since SEOM can visualize ZnO and CuO NPs with high contrast, it could be used to investigate the behavior of NPs in cells without labeling. Thus, we applied SEOM to A549 cells and NPs in those cells.

The A549 cells were cultured on the entrance window in Dulbecco’s modified Eagle’s medium supplemented with 10% fetal bovine serum (DMEM) according to the methods shown in Appendix A. In some experiments the living cells were enclosed in ECs filled with phosphate-buffered saline (PBS) or DMEM without fixing. In other experiments, to make a comparison with conventional phase contrast (PC) images of exactly the same state, the cells were fixed with paraformaldehyde and then encapsulated in ECs filled with PBS. In the present system, the speed of the evacuation and leakage is restricted to very small (15 – 20 min) to prevent damage to the membrane of ECs. Thus, a time lag of typically 30 min is inevitable between the PC imaging in the air and the SEOM imaging in the SEM chamber; during this interval the cell easily changes its shape. Therefore, we had to employ fixation, although it is not essential for SEOM.

Before showing the images of the cells, EB exposure conditions for avoiding cell damages should be discussed. We have clarified the conditions required to prevent electron penetration (Fig. 2). However, heating by the electrons might harm the cells even below the penetration threshold. Conversely, the cells may remain viable for a limited number of penetrating electrons. Therefore, we systematically investigated the damage to the living cells under various EB conditions. To confirm the viability of the cells, the ECs were disassembled after SEOM imaging, and the cells were double-stained with calcein-AM and propidium iodide (PI) and observed by conventional fluorescence microscopy.

We took electron energy E as one parameter, and electron dose D (electrons/nm2) as another because dose is usually discussed as a key factor for damage. As shown in Appendix E, it was found that an EB with E ≤ 1.2 kV and D ≤ 10 electrons/nm2 does not affect the cell viability. As had been anticipated (Fig. 2), the cells are not damaged for values of E below the penetration threshold (0.8 kV for tm = 15 nm and te = 35 nm). In addition, slight penetration up to 1.2 kV was found to be safe. However, if E exceeds 1.5 kV, even a low dose of 1 electron/nm2 can cause fatal damage to the cells. Previously, D has been regarded as the dominant factor in damage [30

30. S. Thiberge, A. Nechushtan, D. Sprinzak, O. Gileadi, V. Behar, O. Zik, Y. Chowers, S. Michaeli, J. Schlessinger, and E. Moses, “Scanning electron microscopy of cells and tissues under fully hydrated conditions,” Proc. Natl. Acad. Sci. U. S. A. 101, 3346–3351 (2004). [CrossRef] [PubMed]

,31

31. N. de Jonge and F. M. Ross, “Electron miocroscopy of specimens in liquid,” Nat. Nanotechnol. 6, 695–704 (2011). [CrossRef] [PubMed]

,33

33. V. E. Cosslett, “Radiation damage in the high resolution electron microscopy of biological materials: a review,” J. Microsc. 113, 113–129 (1978). [CrossRef] [PubMed]

], but E appears to be dominant in the low energy region of E ≃ 1 kV. Since E strongly influences the energy distribution of the transmitted electrons, this is reasonable. Including the results in Fig. 4, all images in this paper were recorded at E ≤ 1.2 kV (mainly E =0.8 – 1.0 kV) and all the cells shown below were observed with D ≤ 10 electrons/nm2 (except for some results in Appendix F).

Images of the cells are exemplified by those shown in Fig. 5. In the CL images, several images are connected in order to cover the whole cell with sufficiently small pixels.

Fig. 5 Images of unstained cells in PBS. (a) CL (cathodoluminescence) image by SEOM system and (b) PC (phase contrast) image of a part of a fixed cell. The upper half of a vertically oriented long cell is shown. (c) Intensity profiles (red: CL, black: PC) between the arrowheads denoted by A in (a) and (b). (d) Raw CL image and (e) PC image of an unstained living cell in PBS. The PC image was recorded 40 min before the CL image. (f) Enhanced CL image spatially bandpass-filtered at 1/150 – 1/34 pixel−1 and (g) major features traced from (f). Observation conditions of the CL images (E, Xp, Td) are (a) 1.0 kV, 37.5 nm, 1 ms and (d) 0.8 kV, 60 nm, 1 ms. Scale bars: 10 μm.

Figures 5(a) and 5(b) show typical images of an unstained fixed cell in PBS. In the CL image [Fig. 5(a)], the cell looks slightly brighter than the surrounding area and cytoplasmic granules are seen as bright spots. Comparison with the PC image of the same area [Fig. 5(b)] shows that only the granules near the edge are visible in the CL image. This suggests that these granules are located within 50 nm of the adhesion surface. In contrast, granules at the center of the cell, visible only in the PC image (denoted by B), are far from the emitting layer. The granules in the CL images look much smaller than those in the PC images, and some are below the diffraction limit [Fig. 5(c)].

Figures 5(d) and 5(e) show unstained living cells in PBS. This cell was confirmed to be alive by double staining after recording this image (see Appendix F). In the CL image [Fig. 5(d)], the entire cell region looks bright and no particular feature can be recognized. Similarly, the PC image [Fig. 5(e)] exhibits no remarkable feature, such as cytoplasmic granules, in this case. However, when properly bandpass-filtered, thin fibers, the outline of the nucleus, and some granules become discernible in the CL image [Figs. 5(f) and 5(g)]. The fibers could be attributed to cytoskelton and all these structures should be located within 50 nm of the adhesion surface.

Next, cells co-incubated with NPs (see Appendix A for methods) were observed. Since ZnO NPs appear bright, they could hardly be distinguished from the inherent cytoplasmic granules that also look bright and we could not obtain any new findings. In contrast, CuO NPs were easily recognized in the cells since they appear dark.

In Figs. 6(a) and 6(b), typical images of a cell containing CuO NPs are shown. In the conventional PC image [Fig. 6(b)], it is difficult to distinguish the cytoplasmic granules and CuO NPs. On the other hand, aggregated CuO NPs can be seen as dark regions in the CL image [Fig. 6(a)]. By use of energy dispersive X-ray spectrometry (EDS), the dark features in the CL images were confirmed as being CuO [Fig. 6(c)]. In Fig. 6(a), the sharpness of the dark region is less than that for the CuO NPs directly deposited on the emitting layer [Fig. 4(d)]. This suggests that the CuO NPs lie slightly away from the adhesion interface but still within the proximity sufficient for energy transfer (≤ 10 nm; i.e., they are located just inside the cell membrane). These features imply that the CuO NPs are taken up into the cells and condensed into several clusters. This is consistent with the aggregated CuO NPs observed in the sliced cells by TEM [46

46. N. Hanagata, F. Zhuang, S. Connolly, J. Li, N. Ogawa, and M. Xu, “Molecular responses of human lung epithelial cells to the toxicity of copper oxide nanoparticles inferred from whole genome expression analysis,”ACS Nano 5, 9326–9338 (2011). [CrossRef] [PubMed]

].

Fig. 6 Typical images of unstained fixed cells in PBS. (a) Raw CL (cathodoluminescence) image by SEOM system and (b) PC (phase contrast) image of a cell that has taken up CuO NPs. (c) EDS (X-ray spectrometry) spectra measured at a dark CL area (upper) and at a blank area (lower). EDS data were recorded for a similar cell. Cu is detected in only the dark areas. In blank areas, O, C, Cl, Na, Al, Si are detected. Al comes from the specimen holder; Cl and Na are from the PBS. (d) Raw CL image and (e) PC image of a cell observed at E = 1.2 kV. (f) CL image spatially bandpass-filtered at 1/100 – 1/75 pixel−1. Both bright cytoplasmic granules (A and B) and dark CuO NPs (C) are visible in (d). However, another feature should be noted here; microstructures of the adhesion surface can be seen even in (d), but are very clear in the filtered image in (f). Cell D in (e) is not seen in (d), because the cell is spherical and in contact with the emitting layer at only a small point. Observation conditions of the CL images (E, Xp, Td) are (a) 1.0 kV, 40 nm, 0.5 ms and (d) 1.2 kV, 60 nm, 1 ms. Scale bars: (a) and (b) 5 μm, (d) – (f) 10 μm.

Figure 6(d) shows images that are interesting from another aspect. This CL image was acquired at the maximum viable energy E = 1.2 kV, which is higher than the penetration threshold. It seems the microstructure of the adhesion surface is visualized as bright areas. For clarity, a spatially bandpass-filtered image is shown in Fig. 6(f). Such features are not seen in the conventional PC image [Fig. 6(e)]. Some of the electrons penetrate the entrance window and hit the cells at this condition. Therefore, the autocathodoluminescence (auto-CL) of the cell might also contribute to the image, and the inhomogeneity in the adhesion state would be sensitively visualized. Since similar structures were frequently observed at E = 1.2 kV, this could be a general feature of A549; but further investigation is necessary.

5. Discussion

The results presented here do not exhibit the final performance of SEOM, as several problems exist in the current setup. The most serious one is the electron durability of the emitting material. Organic materials are easily damaged by the EB; thus, the total number of photons generated from a unit area is limited. This leads to a noisy image, and acquisition of a movie is not practical since the area is bleached by a single scanning.

The second problem is efficiency. The quantum efficiency determined from the relationship between beam current and photon count was 10−5 (see Appendix G). Our system has a potential to acquire images at a rate of 50 frames/s. Nevertheless, due to the low photon emission efficiency in addition to the low current from the cold field-emission gun, we had to set the dwell time as long as 1 ms in order to collect all the producible photons from each pixel; this led to an imaging time of several minutes for one picture.

Another issue is the limited contrast. The image of the current SEOM is based on a small emission enhancement. Incorporation of some new technique for contrast enhancement is crucial. Furthermore, since the image contrast is essentially dependent on the refractive index and the distance from the emitting surface of the object, the interpretation of the image is not necessarily straightforward.

Recently semiconductor superlattices with CL efficiency reaching 40% have been reported [47

47. T. Oto, R. G. Banal, K. Kataoka, M. Funato, and Y. Kawakami, “100 mW deep-ultraviolet emission from aluminium-nitride-based quantum wells pumped by an electron beam,”Nat. Photonics 4, 767–771 (2010). [CrossRef]

]. The uniformity of the emission is of great interest. For improving the efficiency, we can find another clue in Fig. 2(b). In the current configuration, most electrons lose energy in the SiN membrane and only a small fraction contributes to the CL in the emitting layer. If the emitting layer is deposited on the incidence side, the efficiency will be improved drastically but at the cost of sacrificing image resolution, since the distance between the dipole and the specimen will increase. A membrane made of light emitting material would be an ideal solution [48

48. A. Chiba, S. Tanaka, W. Inami, A. Sugita, K. Takada, and Y. Kawata, “Amorphous silicon nitride thin films implanted with cerium ions for cathodoluminescent light source,” Opt. Mater. 35, 1887–1889 (2013). [CrossRef]

].

In Appendix H, an image of a dried cell is shown for reference. Filopodia with a width of 151 nm are clearly observed. This CL image is much sharper than the images in Figs. 5 and 6. Here the greater refractive index contrast led to a higher image contrast and an improved S/N ratio. If a membrane made of an EB-durable, efficient, and nanometrically uniform inorganic material is realized, an image with a S/N ratio as high as that in Appendix H will also be obtained for cells in liquid, due to the increased number of photons. In addition, dynamic observation at a video rate, or even at a higher frame rate, could become possible.

In this study, our primary aim was to demonstrate that SEOM can visualize unstained living cells. However, SEOM should also be applicable to fluorescently labeled specimens like conventional NSOMs. The current green emitting layer would be suitable for exciting red fluorescence. If we use undoped PVK, excitation of green fluorophores would be possible.

Finally, we should address the bursting of the SiN membrane. As one might easily imagine, this does happen. However, the membrane burst during rough evacuation in the airlock before transferring the EC to the main chamber in almost all cases. The membrane always bulges out under vacuum due to the pressure difference, and occasionally fails, presumably owing to the stress caused by the adhered specimen. In addition, leakage of the liquid or the formation of a bubble sometimes happens instead of bursting, due to an imperfect seal at the packing. While we encountered a rupture of the membrane during the imaging only once out of more than 100 trials, there was no damage to the apparatus. The interlock system of the SEM immediately responded to the pressure change and closed a valve to protect the field-emission gun. The vacuum in the chamber recovered within a few seconds, since the amount of the liquid in the EC is limited (4 μl).

6. Conclusion

A near-field scanning optical microscope based on an EC with a cathodoluminescent window has been described. By attaching a detection unit to the specimen stage, an ordinary existing SEM is changed to a super-resolution optical microscope that can non-invasively visualize unstained living cells in liquid. A special inverted SEM is not required. The entrance window of the EC consists of a 20-nm-thick SiN membrane coated with a 30-nm-thick PVK:C6 film and maintains an arbitrary environment filled with liquid or gas. By scanning the surface of the entrance window with a low-energy EB of 0.8 – 1.2 kV, a near-field optical image of the specimen adhered to the opposite surface was acquired without damage to the specimen. The image is formed by enhanced dipole radiation caused by the sample’s higher refractive index or by energy transfer to absorbing materials, and features as small as 62 nm, far beyond the diffraction limit, could be visualized. Furthermore, several observations demonstrated that SEOM has the potential to produce biologically important results. Observations of unstained living human lung epithelial cells in PBS showed fibers and granules near the adhesion interface, which were not resolved by conventional phase microscopy. The cells after the observation were proved to stay alive. In addition, it was shown that the cells took up and aggregated CuO NPs. Undetermined microstructures at the adhesion surface were also revealed.

Stimulated by the success of fluorescent nanoscopy [16

16. S. W. Hell, “Far-field optical nanoscopy,” Science 316, 1153–1158 (2007). [CrossRef] [PubMed]

], development of far-field super-resolution optical microscopy without fluorescent labeling is becoming a hot area. Cotte et al. demonstrated that unstained living biological specimens can be visualized with a resolution of 90 nm by linear wave optics based on digital holographic microscopy and complex deconvolution [17

17. Y. Cotte, F. Toy, P. Jourdain, N. Pavillon, D. Boss, P. Magistretti, P. Marquet, and C. Depeursinge, “Marker-free phase nanoscopy,” Nat. Photonics 7, 113–117 (2013). [CrossRef]

]. Wang et al. observed non-fluorescent graphite nanoplatelets with a resolution of 225 nm by saturated absorption with an intense doughnut-shaped laser beam [18

18. P. Wang, M. N. Slipchenko, J. Mitchell, C. Yang, E. O. Potma, X. Xu, and J. X. Cheng, “Far-field imaging of non-fluorescent species with subdiffraction resolution,” Nat. Photonics 7, 449–453 (2013). [CrossRef]

]. These techniques rely on special optical setup such as opposed oil-immersion objective lenses or femtosecond pump-probe system. In contrast, our method can be applied relatively easily if an SEM is available. We have demonstrated image resolutions higher than that by Wang et al. for both NPs and biological specimens. While our resolution for biological cells is not as high as that by Cotte et al., it should be noted that our results are obtained without any image processing such as deconvolution; if such correction is employed, image quality of SEOM could also be enhanced. However, for practical applications in biology, an improvement in the emitting material is of the first priority.

Appendix A: Materials and methods

Details on the system

The typical beam current, EB diameter under the conditions in this paper, working distance, and chamber pressure are 70 pA, 5 – 10 nm, 6 – 7 mm, and 1×10−4 Pa, respectively. Since our SEM is not equipped with a beam blanker, a mechanical shutter driven by a solenoid was placed below the objective lens so that the entrance window was irradiated with the EB only when the image is acquired. A preamplifier is also placed in the specimen chamber. The whole system is controlled by a graphical interface (LabVIEW, National Instruments). For measuring the CL spectra shown in Fig. 2(c), a light collection unit with a lens and an optical fiber connected to a spectrometer was attached to the specimen stage instead of the PMT detection unit.

Emitting layer

The emitting layer was deposited by micro-injecting a chloroform and o -dichlorobenzene mixed solution of PVK and C6 onto a SiN membrane treated with hexamethyldisilazane. PVK was supplied by Takasago International Corp. C6 was purchased from Tokyo Chemical Industry Co., Ltd. and used as received. The thickness of the PVK:C6 film was evaluated by a Fourier-transform-infrared spectrometer.

Electron scattering simulation

The EB diameter, densities of SiN and PVK, and electron numbers were set at 10 nm, 3.0 g/cm3, 1.2 g/cm3, and 105, respectively [49

49. A. Stoffel, A. Kovács, W. Kronast, and B. Müller, “LPCVD against PECVD for micromechanical applications,” J. Micromech. Microeng. 6, 1–13 (1996). [CrossRef]

, 50

50. J. Han, J. An, R. N. Jana, K. Jung, J. Do, S. Pyo, and C. Im, “Charge carrier photogeneration and hole transport properties of blends of a π-conjugated polymer and an organic-inorganic hybrid material,” Macromol. Res. 17, 894–900 (2009). [CrossRef]

].

Image formation simulation

The refractive indices of water, PS, SiO2, ZnO, and CuO were set at 1.33, 1.59, 1.46, 2.02, and 2.58+i 0.59, respectively [51

51. G. M. Hale and M. R. Querry, “Optical constants of water in the 200-nm to 200-μm wavelength region,” Appl. Opt. 12, 555–563 (1973). [CrossRef] [PubMed]

54

54. K. Postava, H. Sueki, M. Aoyama, T. Yamaguchi, Ch. Ino, Y. Igasaki, and M. Horie, “Spectroscopic ellipsometry of epitaxial ZnO layer on sapphire substrate,” J. Appl. Phys. 87, 7820–7824 (2000). [CrossRef]

]. The SiN membrane and vacuum half space above the sphere were neglected to determine the essential influence of an object on the dipole radiation. Radiation for x, y, and z-directed dipoles were calculated and an incoherent summation of the intensity was considered. For taking account of the electron distribution, we assumed an entrance window with tm = te = 20 nm, a spherical electron range with a radius of 20 nm inscribed in the window, incoherent dipoles uniformly filled in the lower hemisphere, and light collection in the range of θ = 0 – 90°, for simplicity.

Deposition of NPs

The PVK emitting layer of the entrance window was incubated with 0.1% PLL for 30 min and washed four times with water. Then PS (Nisshin EM), SiO2 (Sicaster), ZnO (Sigma-Aldrich), or CuO (Sigma-Aldrich) NPs were suspended in water (concentration: 0.3 – 1 mg/ml), incubated for 1 – 10 min, washed four times, and dried.

Cell culture and co-incubation with NPs

After incubating 0.1% PLL for 30 min and washing it four times with water, a silicone ring was attached around the entrance window [Fig. 1(d)] and 100 μl of A549 cells were seeded with DMEM at a concentration of 5×105 cells/ml. Then the cells were cultured at 37°C in a 5% CO2 humidified incubator for 4 – 48 h. To visualize the viability, the cells were stained using the Cellstain Double Staining Kit (Dojindo). For fixation, the cells were incubated with 4% paraformaldehyde PBS solution for 10 min, and washed with PBS three times. Co-incubation with NPs were carried out as follows; when the cell concentration reached about 70% confluence, the culture medium was replaced with DMEM containing NPs with a concentration of 25 μg/ml [45

45. M. Xu, D. Fujita, S. Kajiwara, T. Minowa, X. Li, T. Takemura, H. Iwai, and N. Hanagata, “Contribution of physicochemical characteristics of nano-oxides to cytotoxicity,” Biomaterials 31, 8022–8031 (2010). [CrossRef] [PubMed]

, 46

46. N. Hanagata, F. Zhuang, S. Connolly, J. Li, N. Ogawa, and M. Xu, “Molecular responses of human lung epithelial cells to the toxicity of copper oxide nanoparticles inferred from whole genome expression analysis,”ACS Nano 5, 9326–9338 (2011). [CrossRef] [PubMed]

] and the cells were cultured for additional 12 –16 h. After washing with PBS twice, the cells were fixed with paraformaldehyde, washed three times, and then enclosed in the ECs with PBS.

Appendix B: Search for emitting layer materials

Before choosing PVK:C6 as the emitting material, various luminescent materials were considered. Although SiN itself exhibits weak CL [Fig. 2(c)], a high current and long dwell time are necessary for obtaining a sufficient number of photons to form an image. Our SiN membranes did not exhibit the ultraviolet CL reported in [35

35. W. Inami, K. Nakajima, A. Miyakawa, and Y. Kawata, “Electron beam excitation assisted optical microscope with ultra-high resolution,” Opt. Express 18, 12897–12902 (2010). [CrossRef] [PubMed]

]. Therefore, emitting layers made of various materials with te = 20 – 60 nm were deposited on SiN membranes with tm = 20 – 100 nm, and spectra and spatial intensity distributions of CL were investigated for EBs with E = 0.8 – 5 kV. Since the space is limited here, the results that led to the choice of PVK:C6 will be simply described.

The SEOM system is a nanometric revival of classical instruments called flying spot scanners, which are based on cathode ray tubes (CRTs). Thus, we first considered the inorganic phosphor materials used for CRTs. As phosphor or inorganic electroluminescent materials with relatively simple compositions, sputtered films of ZnS doped with Ag (0.6 – 1 wt%), ZnO with O vacancies, and ZnS doped with Mn (0.3 – 2 wt%) were examined. All of them required thermal treatment for exhibiting CL, and various annealing conditions were compared. ZnS:Ag appeared to be promising in terms of intensity; however, no satisfactory conditions have been found for any materials in terms of producing the required homogeneity. Figures 7(a)–7(c) show some typical results. The CL showed a spatial distribution at a scale of 50 – 200 nm, which is not acceptable for visualizing intensity distributions caused by the specimens. Established phosphor materials are produced as micrometer-sized particles, so extensive research would be necessary for realizing nanometrically uniform inorganic emitting layers with a thickness on the order of 10 nm. However, we believe that a proper material with an optimum treatment can be found in the future, because the auto-CL of SiN is quite uniform.

Fig. 7 Typical CL intensity distributions for inorganic and organic emitting materials: (a) ZnS:Ag (0.6 wt%) annealed at 1000°C for 0.5 h, (b) ZnS:Ag (0.6 wt%) treated with rapid thermal annealing (RTA) at 800°C for 10 s, (c) ZnO treated with RTA at 700°C for 50 s, and (d) PVK:C6 (1 wt%). The CL image in (d) shows the intensity distribution only due to the shot noise, and is suited to a plain screen for visualizing nanometric features of the specimen. The film thicknesses tm and te, and observation conditions E, Xp, and Td are as follows: (a) 100 nm, 40 nm, 3 kV, 15 nm, 10 ms; (b) 100 nm, 40 nm, 5 kV, 15 nm, 1 ms; (c) 100 nm, 40 nm, 5 kV, 15 nm, 10 ms; and (d) 20 nm, 44 nm, 0.9 kV, 30 nm, 16 ms. Scale bars: 500 nm.

Another type of inorganic material, CdSe/ZnS quantum dots (Sigma-Aldrich, emission peak: 480 nm) embedded in polycarbonate, was also examined. Nevertheless, an emission decrease due to the EB irradiation, which was similar to that for organic materials, was observed. The reason for this unexpected result is not yet clear.

We also investigated various organic materials, such as tris(8-hydroxyquinolinato)-aluminium (Alq, Wako), PVK without C6, plastic scintillator (BC-400, Saint-Gobain), and a few types of polyphenylenevinylene, in addition to PVK:C6. PVK:C6 showed the best performance in terms of intensity and EB durability. As exemplified in Fig. 7(d), the spatial homogeneity of organic materials was generally excellent. Small molecules such as Alq are particularly promising since thermal evaporation is possible. However, they are damaged by water, so they should be deposited on the outer side of the SiN membrane.

Another interesting organic material was the polyimide film of commercial ECs (QX-102, Quantomix). They exhibit strong auto-CL and enable the acquisition of images with a high S/N ratio. Nonetheless, since the commercial films are too thick (≃ 200 nm), the available resolution is limited.

Most of the emitting materials exhibited remarkable nonlinear behavior, presumably due to excitation saturation. Both the spectra and intensities strongly depend on the scanning step (pixel size) and dwell time.

Appendix C: Analysis of emission collection properties based on ray tracing

A ray tracing simulation (TracePro, Lambda Research) was applied to the propagation of rays with a unit intensity emitted in various directions from the center of the membrane, and the fraction of the rays that hit the detector was investigated. The results are shown in Fig. 8. It was found that 21% of the CL emitted into the total solid angle of 4π arrives at the PMT.

Fig. 8 (a) Rays in the xz plane with an intensity higher than 0.2 of that of the original rays are shown for θ = 0, 5, ..., 90°. Right: magnification around the membrane. Emission at high angles is reflected off the sloped surfaces of the Si frame and directed toward the PMT. The coordinate system is shown in the inset of (b). The rays that reach the detector are shown in red, and those do not in blue. Due to the total internal reflection in the exit window, rays at θ ≃ 45° cannot be captured. The shield window on the PMT is a quartz plate coated with an indium tin oxide for shielding the electric field caused by the high voltage applied to the photocathode of the PMT; however, this also contributes to guide the rays to the PMT. (b) The fraction of the power that reaches the PMT compared to rays with a unit intensity emitted in the direction of (ϕ, θ). Right: intensity integrated over ϕ, I(θ), is plotted in the form of I(θ)sinθ. The area surrounded by the curve gives the total detected power.

Appendix D: Influence of refractive index contrast on the image

While simulation results for images of objects in water (n1 = 1.33) were presented in Fig. 3, calculations were also done for particles under a vacuum (n1 = 1). Major results on the intensity profiles for isolated spheres with a diameter 2a = 100 nm are summarized in Fig. 9. Dominant parameters that determine the image are refractive index contrast (n2n1)/n1 and the distance from the emitting layer d. Image contrast (peak height) is proportional to the index contrast, however, the resolution (peak width) stays almost unchanged. Although the image contrast exponentially decays for the distance increase as shown in Fig. 3(c), the resolution is not influenced so much; a particle at d = 50 nm looks slightly larger but still below the diffraction limit. Since the resolution is not sensitive to the index contrast, experimental results for NPs under a vacuum were discussed in Fig. 4.

Fig. 9 Dependence of the image contrast (upper) and peak width (lower) for spheres with a diameter 2a = 100 nm on the refractive index contrast at λ = 550 nm. The values of the index contrast correspond to PS or SiO2 spheres in water or vacuum, as denoted in the lower panel. The particle size 2a = 100 nm is also shown by the horizontal line in the lower panel. Spheres placed on the emitting surface (d = 0 nm) are observed as their actual sizes irrespective of the refractive indices of the sphere and the surrounding medium.

Appendix E: Cell viability under various observation conditions

The results of the double staining of cells observed with various values of energy E and dose D were not necessarily straightforward. Many cells did not show fluorescence for either calcein-AM or PI, even though apparent changes in color and shape were observed. So long as red fluorescence by PI is not observed, these cells cannot be judged to be dead. However, those cells can be regarded as being less viable; thus, we classified such cells as damaged. Some cells were detached from the substrate and disappeared while disassembling the ECs. As shown in Fig. 10, the regions for viable and damaged cells were clearly separated, although a few exceptions can be found. From these results, the parameter ranges of E < 1.2 kV and D < 10 electrons/nm2 were judged as viable conditions for A549 cells, and all of the observations of cells discussed in this paper (except in Appendix F) were carried out within these ranges.

Fig. 10 Viability of 165 cells observed under various EB conditions. The data are plotted with small horizontal shifts so that all the results can be seen. Typical beam current was 45 – 85 pA, and the values of D were adjusted by changing Td in the range of 0.02 – 4 ms. The dose was determined by the measured electron transmittance. The pixel size was Xp = 60 nm. Results for cells filled with both PBS and DMEM are shown in the figure, as there was no difference between them. The cells in this work were observed under EB conditions within the green line.

Appendix F: Typical results of double staining

Typical results used for obtaining Fig. 10 are exemplified by Fig. 11. Figures 11(a)–11(d) show the cell presented in Fig. 5(d), and was confirmed to be living based on the green fluorescence of calcein-AM. Figures 11(e)–11(h) exhibit the results for E = 3 kV. The cell became dark after the acquisition of the CL image, and found to be dead by observing the red fluorescence of PI. The CL and SE images for this cell are also shown in Figs. 11(i) and 11(j) for reference. The CL image is very clear and shows bright and dark spots with sub-diffraction-limit sizes and many other features. In our system, an SE image is simultaneously obtained with the CL image; however, the SE usually visualizes the flat surface of the SiN membrane and shows no particular features. However, in the SE image in Fig. 11(j), structures corresponding to the cell are displayed; this clearly indicates that the EB has penetrated the entrance window and reached the cell. This is equivalent to the images of Thiberge et al. and Nishiyama et al. [30

30. S. Thiberge, A. Nechushtan, D. Sprinzak, O. Gileadi, V. Behar, O. Zik, Y. Chowers, S. Michaeli, J. Schlessinger, and E. Moses, “Scanning electron microscopy of cells and tissues under fully hydrated conditions,” Proc. Natl. Acad. Sci. U. S. A. 101, 3346–3351 (2004). [CrossRef] [PubMed]

, 32

32. H. Nishiyama, M. Suga, T. Ogura, Y. Maruyama, M. Koizumi, K. Mio, S. Kitamura, and C. Sato, “Atmospheric scanning electron microscope observes cells and tissues in open medium through silicon nitride film,” J. Struct. Biol. 169, 438–449 (2010). [CrossRef] [PubMed]

]. Since the cell is directly excited by the electrons, the auto-CL of the cell might also contribute to this CL image. Note that the CL image [Fig. 11(i)] is much clearer and shows more diverse information than the SE image [Fig. 11(j)].

Fig. 11 (a) The PC image after the acquisition of the CL image, (b) that after the double staining, (c) the fluorescence image for calcein-AM, and (d) that for PI. The cell shrank at each stage but it was found to be alive. During the disassembly of the EC and the double staining, another cell nearby was detached and moved into the field of view. (e) – (h) A cell observed using an EB with E = 3 kV and D = 28 electrons/nm2. (e) The PC image before the acquisition of the CL image, (f) that after the CL observation, (g) the fluorescence image for calcein-AM, and (h) that for PI. The fluorescence images were obtained and displayed under the same conditions as (c) and (d). The nucleus of the cell is stained by PI, and this cell is found to be dead. (i) The CL and (j) SE images obtained by EB scanning in the square area in (e). Xp = 60 nm and Td = 1 ms. Scale bars: 10 μm.

Appendix G: Quantum efficiency

Since the number of electrons injected into each pixel is given by the beam current and the dwell time, and the number of the photons is recorded, we can discuss the quantum efficiency, that is, the number of photons per injected electron, of SEOM. Under typical conditions for the results shown in this paper, the quantum efficiency was evaluated to be about 10−5.

As in the discussion by Reimer [3

3. L. Reimer, Scanning Electron Microscopy, 2 (Springer, 1998). [CrossRef]

], the total efficiency of this microscope ηtot can be written as

ηtot=ηel×(E/El)×ηph×ηext×ηcol×ηdet,
(1)

where ηel is the fraction of primary electrons that reaches the emitting layer, El is the mean energy transfer per loss event, ηph is the conversion efficiency from electron-hole (e–h) pairs to photons, ηext is the light extraction efficiency from the high-index emitting layer, ηcol is the yield of the photons extracted from the emitting layer that reach the PMT, and ηdet is the quantum efficiency of the PMT.

Here the electron energy is E ≃ 1 kV, and Cosslett [33

33. V. E. Cosslett, “Radiation damage in the high resolution electron microscopy of biological materials: a review,” J. Microsc. 113, 113–129 (1978). [CrossRef] [PubMed]

] gives El ≃ 10 V. So E/El ≃ 100 e–h pairs are generated per primary electron. From a similar discussion on scintillators by Reimer [3

3. L. Reimer, Scanning Electron Microscopy, 2 (Springer, 1998). [CrossRef]

], the specifications of the PMT, refractive indices, and the discussion in Appendix C, we can assume ηph ≃ 0.03, ηdet ≃ 0.1, ηext ≃ 0.1, and ηcol ≃ 0.2. ηel can be estimated from Fig. 2(b) as ηel ≃ 10−2 – 10−3. Thus, we obtain ηtot = 6× 10−5 – 6×10−6 ≃ 10−5. The good agreement with the experimental result indicates that the assumed efficiency of the emitting layer ηph was reasonable. The discussion here also shows that we still have plenty of room for improvement in ηel.

Appendix H: CL images of dried cells

Figure 12 shows CL images of dried cells that were exposed to the vacuum. Compared with the images of cells in liquid, the images here are very clear. This is because the refractive index of the medium is low which leads to a high index contrast. Filopodia around the cell are clearly visible and some of them have widths narrower than the diffraction limit.

Fig. 12 Typical CL image of dried cells. The square area in (a) is magnified in (b). The profiles between the arrowheads denoted by A and B in (b) are shown in (c). Observation conditions are E = 1.2 kV, Xp = 60 nm, and Td = 1 ms. Scale bars: (a) 10 μm, (b) 5 μm.

Acknowledgments

References and links

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D. W. Pohl, W. Denk, and M. Lanz, “Optical stethoscopy: image recording with resolution λ/20,” Appl. Phys. Lett. 44, 651–653 (1984). [CrossRef]

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A. Lewis, M. Isaacson, A. Harootunian, and A. Muray, “Development of a 500 Å spatial resolution light microscope,” Ultramicroscopy 13, 227–232 (1984). [CrossRef]

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T. Saiki, K. Nishi, and M. Ohtsu, “Low temperature near-field photoluminescence spectroscopy of InGaAs single quantum dots,”Jpn. J. Appl. Phys. 37, 1638–1642 (1998). [CrossRef]

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M. Koopman, A. Cambi, B. I. de Bakker, B. Joosten, C. G. Figdor, N. F. van Hulst, and M. F. Garcia-Parajo, “Near-field scanning optical microscopy in liquid for high resolution single molecule detection on dendritic cells,” FEBS Lett. 573, 6–10 (2004). [CrossRef] [PubMed]

9.

C. Höppener and L. Novotny, “Imaging of membrane proteins using antenna-based optical microscopy,” Nanotechnology 19, 384012 (2008). [CrossRef] [PubMed]

10.

N. Hayazawa, Y. Inouye, Z. Sekkat, and S. Kawata, “Metallized tip amplification of near-field Raman scattering,” Opt. Commun. 183, 333–336 (2000). [CrossRef]

11.

R. Böhme, M. Richter, D. Cialla, P. Rösch, V. Deckert, and J. Popp, “Towards a specific characterisation of components on a cell surface – combined TERS – investigations of lipids and human cells,” J. Raman Spectrosc. 40, 1452–1457 (2009). [CrossRef]

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T. A. Klar, S. Jakobs, M. Dyba, A. Egner, and S. W. Hell, “Fluorescence microscopy with diffraction resolution barrier broken by stimulated emission,” Proc. Natl. Acad. Sci. U. S. A. 97, 8206–8210 (2000). [CrossRef] [PubMed]

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M. G. L. Gustafsson, “Nonlinear structured-illumination microscopy: wide-field fluorescence imaging with theoretically unlimited resolution,” Proc. Natl. Acad. Sci. U. S. A. 102, 13081–13086 (2005). [CrossRef] [PubMed]

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M. J. Rust, M. Bates, and X. Zhuang, “Sub-diffraction-limit imaging by stochastic optical reconstruction microscopy (STORM),” Nat. Methods 3, 793–795 (2006). [CrossRef] [PubMed]

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Y. Cotte, F. Toy, P. Jourdain, N. Pavillon, D. Boss, P. Magistretti, P. Marquet, and C. Depeursinge, “Marker-free phase nanoscopy,” Nat. Photonics 7, 113–117 (2013). [CrossRef]

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E. Kimura, T. Sekiguchi, H. Oikawa, J. Niitsuma, Y. Nakayama, H. Suzuki, M. Kimura, K. Fujii, and T. Ushiki, “Cathodoluminescence imaging for identifying uptaken fluorescence materials in Kupffer cells using scanning electron microscopy,” Arch. Histol. Cytol. 67, 263–270 (2004). [CrossRef] [PubMed]

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J. Niitsuma, H. Oikawa, E. Kimura, T. Ushiki, and T. Sekiguchi, “Cathodoluminescence investigation of organic materials,” J. Electron Microsc. 54, 325–330 (2005). [CrossRef]

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H. Niioka, T. Furukawa, M. Ichimiya, M. Ashida, T. Araki, and M. Hashimoto, “Multicolor cathodoluminescence microscopy for biological imaging with nanophosphors,” Appl. Phys. Express 4, 112402 (2011). [CrossRef]

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30.

S. Thiberge, A. Nechushtan, D. Sprinzak, O. Gileadi, V. Behar, O. Zik, Y. Chowers, S. Michaeli, J. Schlessinger, and E. Moses, “Scanning electron microscopy of cells and tissues under fully hydrated conditions,” Proc. Natl. Acad. Sci. U. S. A. 101, 3346–3351 (2004). [CrossRef] [PubMed]

31.

N. de Jonge and F. M. Ross, “Electron miocroscopy of specimens in liquid,” Nat. Nanotechnol. 6, 695–704 (2011). [CrossRef] [PubMed]

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H. Nishiyama, M. Suga, T. Ogura, Y. Maruyama, M. Koizumi, K. Mio, S. Kitamura, and C. Sato, “Atmospheric scanning electron microscope observes cells and tissues in open medium through silicon nitride film,” J. Struct. Biol. 169, 438–449 (2010). [CrossRef] [PubMed]

33.

V. E. Cosslett, “Radiation damage in the high resolution electron microscopy of biological materials: a review,” J. Microsc. 113, 113–129 (1978). [CrossRef] [PubMed]

34.

D. B. Peckys, P. Mazur, K. L. Gould, and N. de Jonge, “Fully hydrated yeast cells imaged with electron microscopy,” Biophys. J. 100, 2522–2529 (2011). [CrossRef] [PubMed]

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W. Inami, K. Nakajima, A. Miyakawa, and Y. Kawata, “Electron beam excitation assisted optical microscope with ultra-high resolution,” Opt. Express 18, 12897–12902 (2010). [CrossRef] [PubMed]

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Y. Nawa, W. Inami, A. Chiba, A. Ono, A. Miyakawa, Y. Kawata, S. Lin, and S. Terakawa, “Dynamic and high-resolution live cell imaging by direct electron beam excitation,” Opt. Express 20, 5629–5635 (2012). [CrossRef] [PubMed]

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T. Ogura, “Measurement of the unstained biological sample by a novel scanning electron generation X-ray microscope based on SEM,” Biochem. Biophys. Res. Commun. 385, 624–629 (2009). [CrossRef] [PubMed]

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J. Kido, H. Shionoya, and K. Nagai, “Single-layer white light-emitting organic electroluminescent devices based on dye-dispersed poly(N-vinylcarbazole),” Appl. Phys. Lett. 67, 2281–2283 (1995). [CrossRef]

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OCIS Codes
(170.3880) Medical optics and biotechnology : Medical and biological imaging
(180.5810) Microscopy : Scanning microscopy
(250.1500) Optoelectronics : Cathodoluminescence
(180.4243) Microscopy : Near-field microscopy

ToC Category:
Medical Optics and Biotechnology

History
Original Manuscript: August 5, 2013
Revised Manuscript: October 17, 2013
Manuscript Accepted: October 28, 2013
Published: November 8, 2013

Virtual Issues
Vol. 9, Iss. 1 Virtual Journal for Biomedical Optics

Citation
Hideki T. Miyazaki, Takeshi Kasaya, Taro Takemura, Nobutaka Hanagata, Takeshi Yasuda, and Hiroshi Miyazaki, "Diffraction-unlimited optical imaging of unstained living cells in liquid by electron beam scanning of luminescent environmental cells," Opt. Express 21, 28198-28218 (2013)
http://www.opticsinfobase.org/oe/abstract.cfm?URI=oe-21-23-28198


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References

  1. S. Inoué and K. R. Spring, Video Microscopy, the Fundamentals, 2 (Plenum, 1997). [CrossRef]
  2. L. Reimer, Transmission Electron Microscopy, 4 (Springer, 1997). [CrossRef]
  3. L. Reimer, Scanning Electron Microscopy, 2 (Springer, 1998). [CrossRef]
  4. J. E. Mazurkiewicz and P. K. Nakane, “Light and electron microscopic localization of antigens in tissues embedded in polyethylene glycol with a peroxidase-labeled antibody method,” J. Histochem. Cytochem.20, 969–974 (1972). [CrossRef] [PubMed]
  5. D. W. Pohl, W. Denk, and M. Lanz, “Optical stethoscopy: image recording with resolution λ/20,” Appl. Phys. Lett.44, 651–653 (1984). [CrossRef]
  6. A. Lewis, M. Isaacson, A. Harootunian, and A. Muray, “Development of a 500 Å spatial resolution light microscope,” Ultramicroscopy13, 227–232 (1984). [CrossRef]
  7. T. Saiki, K. Nishi, and M. Ohtsu, “Low temperature near-field photoluminescence spectroscopy of InGaAs single quantum dots,”Jpn. J. Appl. Phys.37, 1638–1642 (1998). [CrossRef]
  8. M. Koopman, A. Cambi, B. I. de Bakker, B. Joosten, C. G. Figdor, N. F. van Hulst, and M. F. Garcia-Parajo, “Near-field scanning optical microscopy in liquid for high resolution single molecule detection on dendritic cells,” FEBS Lett.573, 6–10 (2004). [CrossRef] [PubMed]
  9. C. Höppener and L. Novotny, “Imaging of membrane proteins using antenna-based optical microscopy,” Nanotechnology19, 384012 (2008). [CrossRef] [PubMed]
  10. N. Hayazawa, Y. Inouye, Z. Sekkat, and S. Kawata, “Metallized tip amplification of near-field Raman scattering,” Opt. Commun.183, 333–336 (2000). [CrossRef]
  11. R. Böhme, M. Richter, D. Cialla, P. Rösch, V. Deckert, and J. Popp, “Towards a specific characterisation of components on a cell surface – combined TERS – investigations of lipids and human cells,” J. Raman Spectrosc.40, 1452–1457 (2009). [CrossRef]
  12. T. A. Klar, S. Jakobs, M. Dyba, A. Egner, and S. W. Hell, “Fluorescence microscopy with diffraction resolution barrier broken by stimulated emission,” Proc. Natl. Acad. Sci. U. S. A.97, 8206–8210 (2000). [CrossRef] [PubMed]
  13. M. G. L. Gustafsson, “Nonlinear structured-illumination microscopy: wide-field fluorescence imaging with theoretically unlimited resolution,” Proc. Natl. Acad. Sci. U. S. A.102, 13081–13086 (2005). [CrossRef] [PubMed]
  14. E. Betzig, G. H. Patterson, R. Sougrat, O. W. Lindwasser, S. Olenych, J. S. Bonifacino, M. W. Davidson, J. Lippincott-Schwartz, and H. F. Hess, “Imaging intracellular fluorescent protains at nanometer resolution,” Science313, 1642–1645 (2006). [CrossRef] [PubMed]
  15. M. J. Rust, M. Bates, and X. Zhuang, “Sub-diffraction-limit imaging by stochastic optical reconstruction microscopy (STORM),” Nat. Methods3, 793–795 (2006). [CrossRef] [PubMed]
  16. S. W. Hell, “Far-field optical nanoscopy,” Science316, 1153–1158 (2007). [CrossRef] [PubMed]
  17. Y. Cotte, F. Toy, P. Jourdain, N. Pavillon, D. Boss, P. Magistretti, P. Marquet, and C. Depeursinge, “Marker-free phase nanoscopy,” Nat. Photonics7, 113–117 (2013). [CrossRef]
  18. P. Wang, M. N. Slipchenko, J. Mitchell, C. Yang, E. O. Potma, X. Xu, and J. X. Cheng, “Far-field imaging of non-fluorescent species with subdiffraction resolution,” Nat. Photonics7, 449–453 (2013). [CrossRef]
  19. R. Herbst and D. Hoder, “Cathodoluminescence in biological studies,” Scanning1, 35–41 (1978). [CrossRef]
  20. E. Kimura, T. Sekiguchi, H. Oikawa, J. Niitsuma, Y. Nakayama, H. Suzuki, M. Kimura, K. Fujii, and T. Ushiki, “Cathodoluminescence imaging for identifying uptaken fluorescence materials in Kupffer cells using scanning electron microscopy,” Arch. Histol. Cytol.67, 263–270 (2004). [CrossRef] [PubMed]
  21. J. Niitsuma, H. Oikawa, E. Kimura, T. Ushiki, and T. Sekiguchi, “Cathodoluminescence investigation of organic materials,” J. Electron Microsc.54, 325–330 (2005). [CrossRef]
  22. H. Niioka, T. Furukawa, M. Ichimiya, M. Ashida, T. Araki, and M. Hashimoto, “Multicolor cathodoluminescence microscopy for biological imaging with nanophosphors,” Appl. Phys. Express4, 112402 (2011). [CrossRef]
  23. N. Yamamoto, K. Araya, and F. J. García de Abajo, “Photon emission from silver particles induced by a high-energy electron beam,” Phys. Rev. B64, 205419 (2001). [CrossRef]
  24. J. Nelayah, M. Kociak, O. Stéphan, F. J. García de Abajo, M. Tencé, L. Henrard, D. Taverna, I. Pastoriza-Santos, L. M. Liz-Marzán, and C. Colliex, “Mapping surface plasmons on a single metallic nanoparticle,” Nat. Phys.3, 348–353 (2007). [CrossRef]
  25. B. Barwick, D. J. Flannigan, and A. H. Zewail, “Photon-induced near-field electron microscopy,” Nature462, 902–906 (2009). [CrossRef] [PubMed]
  26. A. Kubo, K. Onda, H. Petek, Z. Sun, Y. S. Jung, and H. K. Kim, “Femtosecond imaging of surface plasmon dynamics in a nanostructured silver film,” Nano Lett.5, 1123–1127 (2005). [CrossRef] [PubMed]
  27. D. F. Parsons, V. R. Matricardi, R. C. Moretz, and J. N. Turner, “Electron microscopy and diffraction of wet unstained and unfixed biological objects,” Adv. Biol. Med. Phys.15, 161–270 (1974). [PubMed]
  28. K.-L. Liu, C.-C. Wu, Y.-J. Huang, H.-L. Peng, W.-Y. Chang, P. Chang, L. Hsu, and T.-R. Yew, “Novel microchip for in situ TEM imaging of living organisms and bio-reactions in aqueous conditions,” Lab Chip8, 1915–1921 (2008). [CrossRef] [PubMed]
  29. N. de Jonge, D. B. Peckys, G. J. Kremers, and D. W. Piston, “Electron microscopy of whole cells in liquid with nanometer resolution,” Proc. Natl. Acad. Sci. U. S. A.106, 2159–2164 (2009). [CrossRef] [PubMed]
  30. S. Thiberge, A. Nechushtan, D. Sprinzak, O. Gileadi, V. Behar, O. Zik, Y. Chowers, S. Michaeli, J. Schlessinger, and E. Moses, “Scanning electron microscopy of cells and tissues under fully hydrated conditions,” Proc. Natl. Acad. Sci. U. S. A.101, 3346–3351 (2004). [CrossRef] [PubMed]
  31. N. de Jonge and F. M. Ross, “Electron miocroscopy of specimens in liquid,” Nat. Nanotechnol.6, 695–704 (2011). [CrossRef] [PubMed]
  32. H. Nishiyama, M. Suga, T. Ogura, Y. Maruyama, M. Koizumi, K. Mio, S. Kitamura, and C. Sato, “Atmospheric scanning electron microscope observes cells and tissues in open medium through silicon nitride film,” J. Struct. Biol.169, 438–449 (2010). [CrossRef] [PubMed]
  33. V. E. Cosslett, “Radiation damage in the high resolution electron microscopy of biological materials: a review,” J. Microsc.113, 113–129 (1978). [CrossRef] [PubMed]
  34. D. B. Peckys, P. Mazur, K. L. Gould, and N. de Jonge, “Fully hydrated yeast cells imaged with electron microscopy,” Biophys. J.100, 2522–2529 (2011). [CrossRef] [PubMed]
  35. W. Inami, K. Nakajima, A. Miyakawa, and Y. Kawata, “Electron beam excitation assisted optical microscope with ultra-high resolution,” Opt. Express18, 12897–12902 (2010). [CrossRef] [PubMed]
  36. Y. Nawa, W. Inami, A. Chiba, A. Ono, A. Miyakawa, Y. Kawata, S. Lin, and S. Terakawa, “Dynamic and high-resolution live cell imaging by direct electron beam excitation,” Opt. Express20, 5629–5635 (2012). [CrossRef] [PubMed]
  37. T. Ogura, “Measurement of the unstained biological sample by a novel scanning electron generation X-ray microscope based on SEM,” Biochem. Biophys. Res. Commun.385, 624–629 (2009). [CrossRef] [PubMed]
  38. J. Kido, H. Shionoya, and K. Nagai, “Single-layer white light-emitting organic electroluminescent devices based on dye-dispersed poly(N-vinylcarbazole),” Appl. Phys. Lett.67, 2281–2283 (1995). [CrossRef]
  39. R. R. Lunt, N. C. Giebink, A. A. Belak, J. B. Benziger, and S. R. Forrest, “Exciton diffusion lengths of organic semiconductor thin films measured by spectrally resolved photoluminescence quenching,” J. Appl. Phys.105, 053711 (2009). [CrossRef]
  40. K. Kanaya and S. Okayama, “Penetration and energy-loss theory of electrons in solid targets,” J. Phys. D Appl. Phys.5, 43–58 (1972). [CrossRef]
  41. D. Drouin, A. R. Couture, D. Joly, X. Tastet, V. Aimez, and R. Gauvin, “CASINO V2.42—a fast and easy-to-use modeling tool for scanning electron microscopy and microanalysis users,” Scanning29, 92–101 (2007). [CrossRef] [PubMed]
  42. H. Mertens, A. F. Koenderink, and A. Polman, “Plasmon-enhanced luminescence near noble-metal nanospheres: comparison of exact theory and an improved Gersten and Nitzan model,” Phys. Rev. B76, 115123 (2007). [CrossRef]
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