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Enhanced biological cathodoluminescence
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Optics Communications 281 (2008) 1901–1908 www.elsevier.com/locate/optcom
Enhanced biological cathodoluminescence Phyllis J. Fisher a
a,*
, William S. Wessels a, Allan B. Dietz b, Franklyn G. Prendergast
a
Department of Molecular Pharmacology and Experimental Therapeutics, Division of Transfusion Medicine, and the Mayo Clinic Cancer Center, Mayo Foundation, 200 1st Street SW, Rochester, MN 55905, USA b Department of Laboratory Medicine and Pathology, Division of Transfusion Medicine, and the Mayo Clinic Cancer Center, Mayo Foundation, 200 1st Street SW, Rochester, MN 55905, USA Received 19 January 2007; received in revised form 26 March 2007; accepted 10 April 2007
Abstract We have combined the specificity of antibody labeling, the power of fluorescence detection, and the resolution of scanning electron microscopy (SEM) to identify antigenic sites on nanometer-scale features of mammalian cells. Cathodoluminescence (CL) detection in SEM was used to locate fluorophores bound to antibodies specific for cell surface epitopes. Sample preparation and instrument setup were optimized to yield the maximum luminescence compatible with a high definition secondary electron image. Separable CL component distances of less than 300 nm have been calculated. Antibody-specific fluorophores are associated with unique morphological features on a human dendritic cell. This technology provides a tool to identify the relationship between cell surface structures and receptor-ligand binding or other antigen-defined physiological states. Ó 2007 Elsevier B.V. All rights reserved. Keywords: Scanning electron microscopy; Cathodoluminescence; Fluorescence; Antigen detection; Mammalian cells; Cell morphology; Resolution
1. Introduction The study of sub-cellular processes is dependent upon the relative juxtaposition of the molecules involved, their location within the cell and the limitations imposed by the resolution of the techniques used to study the process. Scanning electron microscopy (SEM) yields high resolution (10–20 nm) [1] of the topological detail of tissues and cells but cannot discern individual proteins or molecules. Confocal microscopy provides highly specific detection of fluorescently tagged molecular labels, but is constrained by the Abbe´ Principle1 which limits the ultimate resolution of this technique to about 200 nm. Neither of these technologies alone can provide the sub-cellular, three-dimensional infor*
Corresponding author. Tel.: +1 507 266 4020; fax: +1 507 284 8111. E-mail address: fi[email protected] (P.J. Fisher). 1 The Abbe´ Principle states that the smallest distance that can be resolved between two lines by optical instruments is proportional to the wavelength and inversely proportional to the angular distribution of the light observed dmin = k/n sin a. 0030-4018/$ - see front matter Ó 2007 Elsevier B.V. All rights reserved. doi:10.1016/j.optcom.2007.04.069
mation necessary to map the location of molecules to identified surface structures of cells. Cathodoluminescence (CL) is light emission resulting from irradiation of luminescent material by an electron beam. In the field emission gun, scanning electron microscope FEG–SEM (Fig. 1A) an electron emission current is produced by an electric field concentrated at the sharp metal tip of the gun cathode. The electron beam is accelerated down the column through a series of electronic lenses and apertures (which are discussed below in the Instrumentation section). The photon collector (Fig. 1B) consists of a focusing mirror set annular to, or at an angle near, the opening of the objective lens. Light emitted by the sample at near-incident angle is collected by this mirror and is piped, optionally through an interference filter or monochromator, to a photomultiplier tube (PMT). When the photon detector is independent of other detectors within the microscope, the CL image can be constructed concurrently with images resulting from other emissions such as secondary electrons (SE).
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Fig. 1. Diagram of the field emission scanning electron microscope. (A) Orientation of the elements of the microscope: the electron beam produced in the electron gun from a sharp tungsten tip with a radius of less than 100 nm, is accelerated through apertures of micron-scale holes in a metal film and electronic lenses both of which control the diameter of the beam, deflector coils which raster the beam over the sample, and the objective lens which focuses the beam on the specimen. (B) The emissive energy collection and data processing layout of this system: Cathodoluminescence emitted from the specimen is collected and focused by a mirror beneath the objective lens. These signals are piped to a photomultiplier where the signals are processed and computer imaged. Secondary electrons are collected in an independent detector and processed in a similar manner.
Cathodoluminescence has been a realized tool in SEM for over forty years [2], but has seen limited use in biological sciences. Many substances both intrinsic (e.g. aromatic amino acids, some nucleic acids, cholesterol and bilirubin) and extrinsic (e.g. whole cell dyes such as acridine orange or antibody-bound fluorescent molecules) will luminesce from electron excitation. In fact, it has been shown that the luminescence properties of macromolecules are independent of the type of exciting radiation [3]. The absorbance spectra of biological materials from electron excitation is similar to those from photon excitation although wavelength shifts may be apparent due to the nonaqueous environment of the electron microscope [4]. However, problems of background energy contamination, sample damage, and detector sensitivity have limited the biological applications of CL. Background contaminations arose from reflected light from the glowing tungsten filament of early electron beam sources, low signal-to-noise level of earlier uncooled PMTs, as well as possible light production from electrons striking contaminant oil or oxide layers on stage components [2]. Sample damage occurred as a result of the necessity of employing high beam energy in order to overcome the high background and low PMT signal-tonoise ratio. As early as 1974, fluorescent amino acids and nucleic acids were detected by CL but a strong signal was observed only in a purified dry powder [5]. Unstained mammalian cells gave off a weak undifferentiated signal [4] while some extrinsic dyes increased the overall luminescence of the cells
or tissues [6,7]. In addition, the instruments and tools available at the time limited image clarity and resolution. Twenty years later the technology had advanced to color differentiation and was applied to the qualitative analysis of bile and cholesterol composition [8,9]. Most recently, specific cells within a tissue, capable of sequestering a circulating fluorescent powder, have been identified by CL [10]. The resolution of the SE image of these cells distinguished specific cells, however the CL areas of the cell were not resolved below a micron, a resolution typical of confocal microscopy. The principle stumbling blocks for advancement in biological SEM–CL have been instrument limitations, quality of sample preparation, and labeling materials. Recent improvements in CL detectors include a mirror (angled or parabolic) annular to the electron beam, a highly reflective or fiber optic light guide, a cooled PMT with a high quantum efficiency (QE) over a wide energy range, and optional monochromator or interference filters. The cooling of the PMT leads to a 102–103 times reduction of dark current [11]. Today there are, broadly available, both primary and secondary fluorescently-labeled antibodies with improved fluorescent labeling efficiencies, and greater photostability [12,13]. Here we describe improvements in sample preparation techniques and instrument parameter adjustments to maximize both the SE signal clarity and the CL signal intensity and resolution, beyond that demonstrated by Kimura et al. [10]. SEM and fluorescence (or CL) detection, combined in a single high resolution scanning event with dual image output, is now a practical tool
P.J. Fisher et al. / Optics Communications 281 (2008) 1901–1908
for identification of molecularly-specific sites on individual biological cells. 2. Sample preparation Three types of samples were used as standards for optimizing the instrument settings. Green fluorescent protein (GFP), isolated from Aequorea forska˚lea [14] and a suspension of Qdot 565 goat anti-mouse IgG antibody conjugate (Invitrogen, Carlsbad, CA) were each applied to aluminum stage mounts and dried under vacuum at room temperature for 12 h. CRT phosphors (red: europium yttrium vanadate, green: zinc cadmium sulfide, and blue: zinc sulfide) were obtained by scraping a cohesive layer from the inside of a computer monitor and transferring these phosphor stripes to carbon tape (SPI, West Chester, PA). Human peripheral blood mononuclear cells, which include T cells, B cells, NK cells and CD14+ monocytes, were isolated from whole blood by centrifugation with Lymphocyte Separation Medium (Organon Teknika, Durham, NC). Several cell preparation steps were developed to improve sample quality and scanning outcomes. We use poly-L-lysine coated (15 min), flat-bottom aluminum weigh dishes as culture plates to aid in the dissipation of excessive charge during electron scanning. After the final sample preparation step, the vertical edge of the aluminum dish is cut down to the bottom surface at several points and the flanges formed are bent down to make a tight contact (ground) with the aluminum microscope stage. Our model for studying cellular morphology is the cells of the monocyte lineage. Monocytes and their derivative cells, dendritic cells, have extensive and diverse morphology. We have found that the extensive, thin, veil-like membrane of dendritic cells is sensitive to environmental changes such as solvent composition or sudden temperature fluctuations. We found that these cells are more structurally stable to fixation if, after removal from a 37 °C incubator, they rest at room temperature for 1 h. Our fixation process begins with 1% paraformaldehyde/PBS (phosphate buffered saline (10 mM phosphate, 137 mM NaCl, 2.7 mM KCl, pH 7.4)) which is added by single drops to a 1:1 volume dilution of the cell medium. Half of this diluted medium is then
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removed by slow pipetting at the edge of the culture plate. These last two steps are repeated three times, thus ending with 1.5% paraformaldehyde. After 30 min the same dilution/pipetting routine is repeated with PBS four times which reduces the residual paraformaldehyde to less than 0.1%. Additional washing steps following label application (below) further reduce the paraformaldehyde concentration to a level that does not interfere with fluorescence. This typical electron microscopy preparation process reduces the loss of loosely attached cells which might be washed off under a stream of buffer. Mouse, anti-human CD14-FITC (fluorescein isothiocyanate) (eBioscience, San Diego, CA) was applied to the sample as per manufacturer’s suggestions, followed by incubation at room temperature for 15 min before washing. A secondary label, goat, anti-mouse DyeMer (488/605) (Invitrogen, Carlsbad, CA) was applied in the same way. (This multiple label was used, at this stage of technique development, only to increase the emission opportunity at each labeled site.) The samples were then subjected to a gradual exchange of aqueous buffer with pure water and then with ethanol followed by critical point drying with liquid CO2 in a pressure/temperature control tank (Ted Pella Inc., Redding, CA). 3. Instrumentation A Hitachi S-4700 FE-SEM (Hitachi, Schaumburg, IL) was used in the dual imaging mode, collecting secondary electrons (SE) on the built-in electron detector and photons on the Centaurus detector (KE Developments Ltd., Cambridge, UK). This detector is in use, at the authors’ institution, as both a backscatter detector (with a scintillator tip) and a CL detector (with an angled mirror tip). As such it is currently fitted with a bialkali type PMT (with a QE greater than 10% in a range from only 320 nm to 530 nm) that is most compatible with the scintillator and less than optimal for cathodoluminescence. This CL detector has a voltage range from 250 V to 1500 V. We usually operate between 750 V and 1000 V. A multialkali type PMT (with a QE greater than 10% in a range of 180– 700 nm) is available and will be useful for multiple fluorophores of different color.
Table 1 Effect of parameter controls on microscope output and the optimal ranges for concurrent CL and SE imaging
Spot size Sharpness Resolution Charging/damage SE signal CL signal Optimal ranges
Accelerating voltage
Beam current
Condenser lens strength
Magnification
Working distance
Higher
Lower
Higher
Lower
Stronger
Weaker
Lower
Higher
Shorter
Longer
# # " " – " 2–10 kV
" # # # – #
" # # " " " 8–15 lA
# " " # # #
# " " " # # 1–6
" # # # " "
– " " # – # 103–104X
– # # " – "
# " " " – # 9–14 mm
" # # # – "
(") indicates and increase in the listed feature. (#) shows a decrease in the listed feature. (–) represents no apparent changes. The last row shows the ranges of parameter controls that were most optimal for CL/SE in our systems.
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The microscope parameter control mechanisms are illustrated in Fig. 1A. Table 1 summarizes the effects of these controls and the optimal parameter ranges for our samples on our system. Both beam accelerating voltage and beam current are controlled at the electron gun. The distance of the stage from the objective lens (working distance) is controlled by the stage goniometer. Magnification is a result of a change in the size of the rastered area of the specimen by the deflection coils. If this size is reduced, when mapped to a constant pixel dimension on the imaging monitor, results in an enlarged image. The electromagnet of the first condenser lens affects the beam probe diameter while stigmatism is controlled by the other condenser lens. The objective lens focuses the beam on the specimen. Operation in the ‘‘Analysis mode’’ of the Hitachi S-4700 enhances CL at the expense of a small loss in resolution. Changes between these ‘‘modes’’ change the lens crossover point thereby affecting spot size and beam current (the specific optical changes have been declared proprietary by company representatives). Digital data from images, used for calculating resolution, was acquired and analyzed with Metamorph software (Molecular Devices Corp., Sunnyvale, CA). Adobe Photoshop was used for image enhancement and cropping. Enhancement was limited to maximizing the pixel spread on the output histogram. No Photoshop filters were used.
Fig. 2. Schematic diagram of the possible energy transitions occurring from the excitation of a fluorophore by an electron beam. E represents the incident beam; S = singlet states; VR = vibrational relaxation; IC = internal conversion; " = absorption; # = emission, Red = X-ray; Blue = secondary electron; Green = photon. Emissions resulting from absorption of the incident beam (A) may propagate photon emission by the reabsorption of emitted secondary electrons (B) and/or X-rays (C). Approximate incident and emission energies are indicated for each transition illustrated. The high energy end for X-ray, of course, could not result from the indicated range of incident energy.
4. Results The production of photons from a fluorescent molecule in the electron microscope may be a complex phenomenon of multiple sources. The beam itself is the primary excitation source but secondary electrons or X-rays produced as a result of energy decay through vibrational relaxation or internal conversion may also act as excitors (Fig. 2). We have schematically illustrated the electronic absorption and X-rays as arising from the third electronic state (S3) however the relative electronic energy separations may be much greater. In fact, the kinetic energy of a single electron may produce multiple photons [15]. The photon output efficiency of the secondary or tertiary sources shown in Fig. 2 is dependent upon the intersection of these deflected
Fig. 3. A scraping of the phosphor layer from a television screen was imaged by secondary electron detection (a) and cathodoluminescence (b, c), with a 450/10 nm bandpass filter (b) and with a 520/10 nm bandpass filter (c). To match the SE image lines to those in the filtered versions we indicate a defect in one of the green lines (arrow) and the computer cursor (cross) on one of the blue lines. The remaining line, with darker edges in the SE image, is the red phosphor. The accelerating voltage was 5 keV, emission current 10 lA, and working distance 18.2 mm. Bar = 500 lm.
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energy vectors with a coulombic field of a target atom. The combined effect of elastic (beam electrons interacting with the nucleus of an atom) and inelastic (beam electrons interacting with the electric field of an atom) events is a redistribution of beam and secondarily produced electrons and photons over an interaction volume. This concept has been illustrated by others in Monte Carlo simulations [16,17]. The interaction volume simulated for a bulk carbon sample extends to over 200 nm below the plane of focus and to over 100 nm laterally from the beam path. Such an interaction volume may be quite different in a biological sample
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but will, in future studies, be considered in a discussion of depth of focus and total emissive volume. Multi-color imaging is possible, as in fluorescence microscopy, and requires only adequate fluorescence intensity and the insertion of appropriate filters or a monochromator. We were able to demonstrate this with the color phosphor lines from a computer monitor. A sample of this coating was imaged with bandpass filters placed between the light-gathering tube and the photomultiplier. The blue phosphors were exposed with a 450/10 nm filter (Fig. 3b) and the green with a 520/10 nm filter (Fig. 3c).
Fig. 4. Antibody-conjugated quantum dots (Qdots) (a, b) were used for calculation of CL resolution. Beam accelerating voltage = 5.0 keV, Z focus = 19.7 mm, magnification = 8.00 k. Bar = 5 lm. SE image (a), CL image showing path (red) of intensity line scan (b), plot of CL intensity from line scan of b (red) and Fourier smoothing of same scan (blue) (c), enlargement of a section of the line scan (d), Fourier analysis of line scan in c (e). Accurate representation of the line scan data requires approximately 3.2 million cycles per meter. This implies useful image components in the 300 nm range. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
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The goal of CL resolution in our system is the identification of the location of fluorescent tags (102–1 lM) on biological cells (5–20 lM). To define the resolution of our system we have chosen to apply the method of Ira Rosenberg particularly because it is operator-independent; that is, free of the prejudice of the analyst in deciding which peaks to measure, calculating local minima and maxima, and deciding on the limit of noise [18]. Quantum dots (Invitrogen, Corp., Carlsbad, CA, USA) (Fig. 4) were used as a standard for this calculation since they are a photostable fluorophore which is becoming more commonly used in fluorescence microscopy as a conjugate on immunoprobes. Rosenberg’s data processing approach includes using fast Fourier transforms on multiple line samples allowing frequency specific averaging, followed by extrapolated smoothing through repeated applications of inverse transformation and transformation. Signal processing limitations were a function of original image data intensity integer values between 0 and 255. Intensity levels from lines drawn through a CL image of a Qdot aggregate provided the digital edge effect used for this calculation. The resultant graph of the Fourier analysis of the line scan data (Fig. 4c) is displayed in Fig. 4d. From averaging three such line scans we find that approximately 3.2 million cycles per meter accurately represents the line scan data implying (by inversion) useful image components in the 300 nm range. To determine if commonly used extrinsic organic fluorescent labels would produce CL in our system we tested fluorescein, rhodamine and green fluorescent protein (GFP). The first two powdered substances (Sigma–Aldrich, St. Louis, MO, USA), commercially obtained, had relatively (compared to GFP) level CL emission across an image field. In the GFP sample bright fluorescence was emitted from select particles while other particles of similar structure in the SE image were not detected in CL (Fig. 5). This GFP, isolated from the jelly fish Aequorea Forska˚lea, had not been taken through final steps of purification. It is interesting that non-CL emitting substances in the sample are isolated as quasi-crystals rather than mixed within the crystal-like particles of emitting protein. Such non-emitting particles may be aggregates of mis-folded or degraded proteins. This topic is not pursued in this study but suggests a new analytical tool for protein structure studies. Cloned GFP is now available in a purer state and will, undoubtedly be used in future biological CL studies. Although the steps taken to electrically ground samples (direct attachment to either an aluminum stage or carbon tape attached to the stage) were adequate for the standard samples employed here, this was not the case for biological cells. Human blood cells were mounted to aluminum plates with a very thin coating of poly-L-lysine but, despite reducing the accelerating voltage to 2 keV and increasing the image acquisition rate from 160 s to 80 s, an electron charge (detectable as a flat white irregular area on the image resulting from detector saturation) obscured sample structure. In addition, the structure of the cells was altered after several such scans (this was determined by viewing the
Fig. 5. SE (a), CL (b), and overlay (c) images of a lyophilized powder of GFP (green fluorescent protein). Beam accelerating voltage = 10 keV, Z focus = 19 mm, magnification = 3.6 K. Bar = 10 lm.
same sample following gold sputter coating where these damaged areas of the sample could be identified in relation to adjacent undamaged cells which had not been irradiated). We tested fluorescent/nanogold dual-labeled probes (Fab’ fragments of polyclonal IgG covalently conjugated to a fluorescent dye molecule and a 1.4 nm gold particle) (Invitrogen, Carlsbad, CA) as a possible discharge mechanism with controlled, even and minimal distribution of the gold by necessarily binding to the antibody targets on the cell. This resulted in no reduction of charging but, in addition, we noted no loss of fluorescence such as reported by others in the use of gold coatings [2,3,19,20]. Therefore we tested osmium and gold coatings of various concentrations. Osmium provided little reduction of sample charging even to the degree that fluorescence was almost totally obscured. However, a 15 s sputter coating of gold–palladium (15 mA, 2.2 kV, using Argon plasma)
P.J. Fisher et al. / Optics Communications 281 (2008) 1901–1908
(Bio-Rad, E-5400, Hercules, CA) greatly reduced the incidence of charging on the cells while passing about 50% of the emission observed without coating. A 60 s coating at the same power settings obliterated detectable fluorescence. Fo¨rster quenching distance calculations [20,21] show that a 12 nm or greater separation between a fluorophore and a gold particle prevents energy transfer/quenching. The continuity of gold particle deposition as well as the overall concentration and depth resulting from different power settings and sputter times should be considered in the determination of the most efficient level of gold deposition. Monocytes are a subset of mononuclear cells found in circulating blood. They are distinguished from other peripheral mononuclear cells (PBMNC) by the presence of a cell surface membrane-associated protein called CD14. Monocytes are also unique in that they possess extensive surface membrane which forms ‘‘veil-like’’ pro-
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trusions. Other PBMNC are more spherical and have short protrusions. A fluorescently tagged antibody to the CD14 protein, incubated with a sample of PBMNC, selectively labels the c cells and is routinely used in fluorescence activated cell scanning (FACS) to identify and select this population. CL detection could be used to grossly identify the CD14+ cells (Fig. 6b) as well as identify typical monocyte morphology and the distribution of surface proteins on the cells. Fig. 6 shows these morphological differences and the CL identification of the CD14+ cells. The fluorescence collected (b) from a select few of the cells in the field of Fig. 6a shows that the power of specific labeling, which is used routinely in confocal microscopy, is translatable to SEM-CL. As demonstrated in (c–f), regions bearing a high concentration of label can be associated with some SE differentiated structures such as protrusions (f, arrow) and veil edges.
Fig. 6. Cathodoluminescence images of fluorescent labels attached to human peripheral blood mononuclear cells. The cells were labeled with anti-human CD14-FITC/Alexa 656 and lightly sputter coated with gold after critical point drying. SE images (a, c), CL images (b, d). Images (a and b) are a field of cells of which only four have the veil-like shape and approximate size of CD14+ cells and display cathodoluminescence. Images (c and d) are magnifications of a similarly cathodoluminescent cell in this sample. Image (e) is an overlay of (d) on (c) and (f) is an enlargement of the boxed area in (e). Higher concentrations of the label occur on the edges of the veils and on some specific structures (arrow). Beam accelerating voltage = 5 keV, (a, b) Bar = 10 lm, (c–e) Bar = 600 nm, (f) Bar = 200 nm.
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5. Conclusion No effective method has previously been demonstrated to link single, or a combination of potentially interacting, molecules to SEM resolved cellular structures. We have combined CL with SE detection to image specific antibody labels on biological cells at the submicron scale. This will be particularly useful in visualizing tagged molecules on specific cell structures, such as veils, dendrites, or gap junctions, where a lowered concentration, or the absence, of a population of receptors or ligands might evince a pathological condition. During the optimization experiments we found that the best CL images were obtained with the highest current range of the electron probe, low beam acceleration voltages (3–5 keV) and a beam current near 10 lA. In addition, we found it necessary for samples to be well grounded to avoid charge buildup. Plating cells with a thin adherent material on an aluminum surface that can be firmly attached to the instrument stage aids in this precaution. An overcoat of gold of limited concentration can help in charge dissipation, but this must be monitored against a loss of fluorescence due to quenching. Fixatives must be thoroughly washed out before labeling to avoid chemical interactions and glutaraldehyde should be avoided due to its contribution to autofluorescence. The effectiveness of SEM-CL as a tool for studying biological samples can be improved. We must explore the relationship between accelerating voltage and depth of electron penetration and the effect of electron scattering on the size of the excitation volume. The emissive volume and its relationship to secondary or tertiary energy sources must also be measured. These aspects will more accurately locate the three-dimensional origin of a photonic signal. In addition, simultaneous detection of labels of multiple colors is a necessity in studies of biological interactions. An enhanced SEM-CL optimized for biological applications would incorporate a PMT of the highest sensitivity and widest spectral range available such as the Hamamatsu R928P which has, a single emission gathering umbrella sensitive to all the emission wavelengths and covering a maximum of the sample surface, simultaneous detection of X-rays, back-scattered electrons, secondary electrons and photons, and separation of the photonic signal by color. Such instrumentation, in combination with recently developed highly photostabile fluorophores and quantum dots would increase the opportunity of identifying molecular components of nanoscale morphological structures on
cells and tissues and their cellular distribution relative to each other. Acknowledgement This investigation was supported, in part, by Grants NCI P30 CA 15083, NIH P50 CA 108961, NCI P50 CA 91956-6 and by Mayo Foundation. We also thank Dr. Angela Klaus, Director of the Microscopy and Imaging Facility, American Museum of Natural History, New York, for support and encouragement. References [1] C.E. Norman, Microscopy and Analysis 53 (2002) 13. [2] E.F. Bond, D. Beresford, G.H. Haggis, Journal of Microscopy 100 (1974) 271. [3] W.A. Barnett, M.L. Wise, E.C. Jones, Journal of Microscopy 105 (1975) 299. [4] P.V.C. Hough, W.R. McKinney, M.C. Ledbetter, et al., Proceedings of the National Academy of Sciences of the United States of America 73 (1976) 317. [5] M.D. Muir, P.R. Grant (Eds.), Cathodoluminescence, Academic Press, London, 1974. [6] W. Brocker, G. Hauck, R. Blaschke, et al., Microscopica Acta Supplement 2 (1978) 260. [7] W. Brocker, G. Pfefferkorn, Scanning Electron Microscopy 2 (1979) 125. [8] A.S. Loginov, S.M. Chebanov, G.V. Saparin, et al., Scanning 20 (1998) 442. [9] G. Ning, T. Fujimoto, H. Koike, et al., Journal of Histochemistry and Cytochemistry 41 (1993) 617. [10] E. Kimura, T. Sekiguchi, H. Oikawa, Archives of Histology and Cytology 67 (2004) 263. [11] M. Cole, D. Ryer, Electro Optical Systems Design, Milton S. Kiver Publications Inc., 1972, pp. 16. [12] W.C. Chan, S. Nie, Science 281 (1998) 2016. [13] N. Panchuk–Voloshina, R.P. Haugland, J. Bishop–Stewart, et al., Journal of Histochemistry and Cytochemistry 47 (1999) 1179. [14] F.G. Prendergast, K.G. Mann, Biochemistry 17 (1978) 3448. [15] A. Einstein, Annual Physik 17 (1905) 132. [16] B. Hafner, Scanning Electron Microscopy Primer, University of Minnesota, Characterization Facility (2007). Available from:. [17] E. Napchan, D.B. Holt, Institute of Physics Conference Series 87 (1987) 733. [18] I. Rosenberg, Proceedings of the SPIE 5375 (2004) 244. [19] P.P.H. Cheng, D. Silvester, G. Wang, et al., J. Phys. Chem. B 110 (2006) 4637. [20] R.D. Powell, C.M.R. Halsey, E. Gutierrez, et al., Proceedings of the National Academy of Sciences of the United States of America (1998) 992. [21] R.D. Powell, C.M. Halsey, J.F. Hainfeld, Microscopy Research and Technique 42 (1998) 2.
Optics Communications 281 (2008) 1901–1908 www.elsevier.com/locate/optcom
Enhanced biological cathodoluminescence Phyllis J. Fisher a
a,*
, William S. Wessels a, Allan B. Dietz b, Franklyn G. Prendergast
a
Department of Molecular Pharmacology and Experimental Therapeutics, Division of Transfusion Medicine, and the Mayo Clinic Cancer Center, Mayo Foundation, 200 1st Street SW, Rochester, MN 55905, USA b Department of Laboratory Medicine and Pathology, Division of Transfusion Medicine, and the Mayo Clinic Cancer Center, Mayo Foundation, 200 1st Street SW, Rochester, MN 55905, USA Received 19 January 2007; received in revised form 26 March 2007; accepted 10 April 2007
Abstract We have combined the specificity of antibody labeling, the power of fluorescence detection, and the resolution of scanning electron microscopy (SEM) to identify antigenic sites on nanometer-scale features of mammalian cells. Cathodoluminescence (CL) detection in SEM was used to locate fluorophores bound to antibodies specific for cell surface epitopes. Sample preparation and instrument setup were optimized to yield the maximum luminescence compatible with a high definition secondary electron image. Separable CL component distances of less than 300 nm have been calculated. Antibody-specific fluorophores are associated with unique morphological features on a human dendritic cell. This technology provides a tool to identify the relationship between cell surface structures and receptor-ligand binding or other antigen-defined physiological states. Ó 2007 Elsevier B.V. All rights reserved. Keywords: Scanning electron microscopy; Cathodoluminescence; Fluorescence; Antigen detection; Mammalian cells; Cell morphology; Resolution
1. Introduction The study of sub-cellular processes is dependent upon the relative juxtaposition of the molecules involved, their location within the cell and the limitations imposed by the resolution of the techniques used to study the process. Scanning electron microscopy (SEM) yields high resolution (10–20 nm) [1] of the topological detail of tissues and cells but cannot discern individual proteins or molecules. Confocal microscopy provides highly specific detection of fluorescently tagged molecular labels, but is constrained by the Abbe´ Principle1 which limits the ultimate resolution of this technique to about 200 nm. Neither of these technologies alone can provide the sub-cellular, three-dimensional infor*
Corresponding author. Tel.: +1 507 266 4020; fax: +1 507 284 8111. E-mail address: fi[email protected] (P.J. Fisher). 1 The Abbe´ Principle states that the smallest distance that can be resolved between two lines by optical instruments is proportional to the wavelength and inversely proportional to the angular distribution of the light observed dmin = k/n sin a. 0030-4018/$ - see front matter Ó 2007 Elsevier B.V. All rights reserved. doi:10.1016/j.optcom.2007.04.069
mation necessary to map the location of molecules to identified surface structures of cells. Cathodoluminescence (CL) is light emission resulting from irradiation of luminescent material by an electron beam. In the field emission gun, scanning electron microscope FEG–SEM (Fig. 1A) an electron emission current is produced by an electric field concentrated at the sharp metal tip of the gun cathode. The electron beam is accelerated down the column through a series of electronic lenses and apertures (which are discussed below in the Instrumentation section). The photon collector (Fig. 1B) consists of a focusing mirror set annular to, or at an angle near, the opening of the objective lens. Light emitted by the sample at near-incident angle is collected by this mirror and is piped, optionally through an interference filter or monochromator, to a photomultiplier tube (PMT). When the photon detector is independent of other detectors within the microscope, the CL image can be constructed concurrently with images resulting from other emissions such as secondary electrons (SE).
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Fig. 1. Diagram of the field emission scanning electron microscope. (A) Orientation of the elements of the microscope: the electron beam produced in the electron gun from a sharp tungsten tip with a radius of less than 100 nm, is accelerated through apertures of micron-scale holes in a metal film and electronic lenses both of which control the diameter of the beam, deflector coils which raster the beam over the sample, and the objective lens which focuses the beam on the specimen. (B) The emissive energy collection and data processing layout of this system: Cathodoluminescence emitted from the specimen is collected and focused by a mirror beneath the objective lens. These signals are piped to a photomultiplier where the signals are processed and computer imaged. Secondary electrons are collected in an independent detector and processed in a similar manner.
Cathodoluminescence has been a realized tool in SEM for over forty years [2], but has seen limited use in biological sciences. Many substances both intrinsic (e.g. aromatic amino acids, some nucleic acids, cholesterol and bilirubin) and extrinsic (e.g. whole cell dyes such as acridine orange or antibody-bound fluorescent molecules) will luminesce from electron excitation. In fact, it has been shown that the luminescence properties of macromolecules are independent of the type of exciting radiation [3]. The absorbance spectra of biological materials from electron excitation is similar to those from photon excitation although wavelength shifts may be apparent due to the nonaqueous environment of the electron microscope [4]. However, problems of background energy contamination, sample damage, and detector sensitivity have limited the biological applications of CL. Background contaminations arose from reflected light from the glowing tungsten filament of early electron beam sources, low signal-to-noise level of earlier uncooled PMTs, as well as possible light production from electrons striking contaminant oil or oxide layers on stage components [2]. Sample damage occurred as a result of the necessity of employing high beam energy in order to overcome the high background and low PMT signal-tonoise ratio. As early as 1974, fluorescent amino acids and nucleic acids were detected by CL but a strong signal was observed only in a purified dry powder [5]. Unstained mammalian cells gave off a weak undifferentiated signal [4] while some extrinsic dyes increased the overall luminescence of the cells
or tissues [6,7]. In addition, the instruments and tools available at the time limited image clarity and resolution. Twenty years later the technology had advanced to color differentiation and was applied to the qualitative analysis of bile and cholesterol composition [8,9]. Most recently, specific cells within a tissue, capable of sequestering a circulating fluorescent powder, have been identified by CL [10]. The resolution of the SE image of these cells distinguished specific cells, however the CL areas of the cell were not resolved below a micron, a resolution typical of confocal microscopy. The principle stumbling blocks for advancement in biological SEM–CL have been instrument limitations, quality of sample preparation, and labeling materials. Recent improvements in CL detectors include a mirror (angled or parabolic) annular to the electron beam, a highly reflective or fiber optic light guide, a cooled PMT with a high quantum efficiency (QE) over a wide energy range, and optional monochromator or interference filters. The cooling of the PMT leads to a 102–103 times reduction of dark current [11]. Today there are, broadly available, both primary and secondary fluorescently-labeled antibodies with improved fluorescent labeling efficiencies, and greater photostability [12,13]. Here we describe improvements in sample preparation techniques and instrument parameter adjustments to maximize both the SE signal clarity and the CL signal intensity and resolution, beyond that demonstrated by Kimura et al. [10]. SEM and fluorescence (or CL) detection, combined in a single high resolution scanning event with dual image output, is now a practical tool
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for identification of molecularly-specific sites on individual biological cells. 2. Sample preparation Three types of samples were used as standards for optimizing the instrument settings. Green fluorescent protein (GFP), isolated from Aequorea forska˚lea [14] and a suspension of Qdot 565 goat anti-mouse IgG antibody conjugate (Invitrogen, Carlsbad, CA) were each applied to aluminum stage mounts and dried under vacuum at room temperature for 12 h. CRT phosphors (red: europium yttrium vanadate, green: zinc cadmium sulfide, and blue: zinc sulfide) were obtained by scraping a cohesive layer from the inside of a computer monitor and transferring these phosphor stripes to carbon tape (SPI, West Chester, PA). Human peripheral blood mononuclear cells, which include T cells, B cells, NK cells and CD14+ monocytes, were isolated from whole blood by centrifugation with Lymphocyte Separation Medium (Organon Teknika, Durham, NC). Several cell preparation steps were developed to improve sample quality and scanning outcomes. We use poly-L-lysine coated (15 min), flat-bottom aluminum weigh dishes as culture plates to aid in the dissipation of excessive charge during electron scanning. After the final sample preparation step, the vertical edge of the aluminum dish is cut down to the bottom surface at several points and the flanges formed are bent down to make a tight contact (ground) with the aluminum microscope stage. Our model for studying cellular morphology is the cells of the monocyte lineage. Monocytes and their derivative cells, dendritic cells, have extensive and diverse morphology. We have found that the extensive, thin, veil-like membrane of dendritic cells is sensitive to environmental changes such as solvent composition or sudden temperature fluctuations. We found that these cells are more structurally stable to fixation if, after removal from a 37 °C incubator, they rest at room temperature for 1 h. Our fixation process begins with 1% paraformaldehyde/PBS (phosphate buffered saline (10 mM phosphate, 137 mM NaCl, 2.7 mM KCl, pH 7.4)) which is added by single drops to a 1:1 volume dilution of the cell medium. Half of this diluted medium is then
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removed by slow pipetting at the edge of the culture plate. These last two steps are repeated three times, thus ending with 1.5% paraformaldehyde. After 30 min the same dilution/pipetting routine is repeated with PBS four times which reduces the residual paraformaldehyde to less than 0.1%. Additional washing steps following label application (below) further reduce the paraformaldehyde concentration to a level that does not interfere with fluorescence. This typical electron microscopy preparation process reduces the loss of loosely attached cells which might be washed off under a stream of buffer. Mouse, anti-human CD14-FITC (fluorescein isothiocyanate) (eBioscience, San Diego, CA) was applied to the sample as per manufacturer’s suggestions, followed by incubation at room temperature for 15 min before washing. A secondary label, goat, anti-mouse DyeMer (488/605) (Invitrogen, Carlsbad, CA) was applied in the same way. (This multiple label was used, at this stage of technique development, only to increase the emission opportunity at each labeled site.) The samples were then subjected to a gradual exchange of aqueous buffer with pure water and then with ethanol followed by critical point drying with liquid CO2 in a pressure/temperature control tank (Ted Pella Inc., Redding, CA). 3. Instrumentation A Hitachi S-4700 FE-SEM (Hitachi, Schaumburg, IL) was used in the dual imaging mode, collecting secondary electrons (SE) on the built-in electron detector and photons on the Centaurus detector (KE Developments Ltd., Cambridge, UK). This detector is in use, at the authors’ institution, as both a backscatter detector (with a scintillator tip) and a CL detector (with an angled mirror tip). As such it is currently fitted with a bialkali type PMT (with a QE greater than 10% in a range from only 320 nm to 530 nm) that is most compatible with the scintillator and less than optimal for cathodoluminescence. This CL detector has a voltage range from 250 V to 1500 V. We usually operate between 750 V and 1000 V. A multialkali type PMT (with a QE greater than 10% in a range of 180– 700 nm) is available and will be useful for multiple fluorophores of different color.
Table 1 Effect of parameter controls on microscope output and the optimal ranges for concurrent CL and SE imaging
Spot size Sharpness Resolution Charging/damage SE signal CL signal Optimal ranges
Accelerating voltage
Beam current
Condenser lens strength
Magnification
Working distance
Higher
Lower
Higher
Lower
Stronger
Weaker
Lower
Higher
Shorter
Longer
# # " " – " 2–10 kV
" # # # – #
" # # " " " 8–15 lA
# " " # # #
# " " " # # 1–6
" # # # " "
– " " # – # 103–104X
– # # " – "
# " " " – # 9–14 mm
" # # # – "
(") indicates and increase in the listed feature. (#) shows a decrease in the listed feature. (–) represents no apparent changes. The last row shows the ranges of parameter controls that were most optimal for CL/SE in our systems.
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The microscope parameter control mechanisms are illustrated in Fig. 1A. Table 1 summarizes the effects of these controls and the optimal parameter ranges for our samples on our system. Both beam accelerating voltage and beam current are controlled at the electron gun. The distance of the stage from the objective lens (working distance) is controlled by the stage goniometer. Magnification is a result of a change in the size of the rastered area of the specimen by the deflection coils. If this size is reduced, when mapped to a constant pixel dimension on the imaging monitor, results in an enlarged image. The electromagnet of the first condenser lens affects the beam probe diameter while stigmatism is controlled by the other condenser lens. The objective lens focuses the beam on the specimen. Operation in the ‘‘Analysis mode’’ of the Hitachi S-4700 enhances CL at the expense of a small loss in resolution. Changes between these ‘‘modes’’ change the lens crossover point thereby affecting spot size and beam current (the specific optical changes have been declared proprietary by company representatives). Digital data from images, used for calculating resolution, was acquired and analyzed with Metamorph software (Molecular Devices Corp., Sunnyvale, CA). Adobe Photoshop was used for image enhancement and cropping. Enhancement was limited to maximizing the pixel spread on the output histogram. No Photoshop filters were used.
Fig. 2. Schematic diagram of the possible energy transitions occurring from the excitation of a fluorophore by an electron beam. E represents the incident beam; S = singlet states; VR = vibrational relaxation; IC = internal conversion; " = absorption; # = emission, Red = X-ray; Blue = secondary electron; Green = photon. Emissions resulting from absorption of the incident beam (A) may propagate photon emission by the reabsorption of emitted secondary electrons (B) and/or X-rays (C). Approximate incident and emission energies are indicated for each transition illustrated. The high energy end for X-ray, of course, could not result from the indicated range of incident energy.
4. Results The production of photons from a fluorescent molecule in the electron microscope may be a complex phenomenon of multiple sources. The beam itself is the primary excitation source but secondary electrons or X-rays produced as a result of energy decay through vibrational relaxation or internal conversion may also act as excitors (Fig. 2). We have schematically illustrated the electronic absorption and X-rays as arising from the third electronic state (S3) however the relative electronic energy separations may be much greater. In fact, the kinetic energy of a single electron may produce multiple photons [15]. The photon output efficiency of the secondary or tertiary sources shown in Fig. 2 is dependent upon the intersection of these deflected
Fig. 3. A scraping of the phosphor layer from a television screen was imaged by secondary electron detection (a) and cathodoluminescence (b, c), with a 450/10 nm bandpass filter (b) and with a 520/10 nm bandpass filter (c). To match the SE image lines to those in the filtered versions we indicate a defect in one of the green lines (arrow) and the computer cursor (cross) on one of the blue lines. The remaining line, with darker edges in the SE image, is the red phosphor. The accelerating voltage was 5 keV, emission current 10 lA, and working distance 18.2 mm. Bar = 500 lm.
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energy vectors with a coulombic field of a target atom. The combined effect of elastic (beam electrons interacting with the nucleus of an atom) and inelastic (beam electrons interacting with the electric field of an atom) events is a redistribution of beam and secondarily produced electrons and photons over an interaction volume. This concept has been illustrated by others in Monte Carlo simulations [16,17]. The interaction volume simulated for a bulk carbon sample extends to over 200 nm below the plane of focus and to over 100 nm laterally from the beam path. Such an interaction volume may be quite different in a biological sample
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but will, in future studies, be considered in a discussion of depth of focus and total emissive volume. Multi-color imaging is possible, as in fluorescence microscopy, and requires only adequate fluorescence intensity and the insertion of appropriate filters or a monochromator. We were able to demonstrate this with the color phosphor lines from a computer monitor. A sample of this coating was imaged with bandpass filters placed between the light-gathering tube and the photomultiplier. The blue phosphors were exposed with a 450/10 nm filter (Fig. 3b) and the green with a 520/10 nm filter (Fig. 3c).
Fig. 4. Antibody-conjugated quantum dots (Qdots) (a, b) were used for calculation of CL resolution. Beam accelerating voltage = 5.0 keV, Z focus = 19.7 mm, magnification = 8.00 k. Bar = 5 lm. SE image (a), CL image showing path (red) of intensity line scan (b), plot of CL intensity from line scan of b (red) and Fourier smoothing of same scan (blue) (c), enlargement of a section of the line scan (d), Fourier analysis of line scan in c (e). Accurate representation of the line scan data requires approximately 3.2 million cycles per meter. This implies useful image components in the 300 nm range. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
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The goal of CL resolution in our system is the identification of the location of fluorescent tags (102–1 lM) on biological cells (5–20 lM). To define the resolution of our system we have chosen to apply the method of Ira Rosenberg particularly because it is operator-independent; that is, free of the prejudice of the analyst in deciding which peaks to measure, calculating local minima and maxima, and deciding on the limit of noise [18]. Quantum dots (Invitrogen, Corp., Carlsbad, CA, USA) (Fig. 4) were used as a standard for this calculation since they are a photostable fluorophore which is becoming more commonly used in fluorescence microscopy as a conjugate on immunoprobes. Rosenberg’s data processing approach includes using fast Fourier transforms on multiple line samples allowing frequency specific averaging, followed by extrapolated smoothing through repeated applications of inverse transformation and transformation. Signal processing limitations were a function of original image data intensity integer values between 0 and 255. Intensity levels from lines drawn through a CL image of a Qdot aggregate provided the digital edge effect used for this calculation. The resultant graph of the Fourier analysis of the line scan data (Fig. 4c) is displayed in Fig. 4d. From averaging three such line scans we find that approximately 3.2 million cycles per meter accurately represents the line scan data implying (by inversion) useful image components in the 300 nm range. To determine if commonly used extrinsic organic fluorescent labels would produce CL in our system we tested fluorescein, rhodamine and green fluorescent protein (GFP). The first two powdered substances (Sigma–Aldrich, St. Louis, MO, USA), commercially obtained, had relatively (compared to GFP) level CL emission across an image field. In the GFP sample bright fluorescence was emitted from select particles while other particles of similar structure in the SE image were not detected in CL (Fig. 5). This GFP, isolated from the jelly fish Aequorea Forska˚lea, had not been taken through final steps of purification. It is interesting that non-CL emitting substances in the sample are isolated as quasi-crystals rather than mixed within the crystal-like particles of emitting protein. Such non-emitting particles may be aggregates of mis-folded or degraded proteins. This topic is not pursued in this study but suggests a new analytical tool for protein structure studies. Cloned GFP is now available in a purer state and will, undoubtedly be used in future biological CL studies. Although the steps taken to electrically ground samples (direct attachment to either an aluminum stage or carbon tape attached to the stage) were adequate for the standard samples employed here, this was not the case for biological cells. Human blood cells were mounted to aluminum plates with a very thin coating of poly-L-lysine but, despite reducing the accelerating voltage to 2 keV and increasing the image acquisition rate from 160 s to 80 s, an electron charge (detectable as a flat white irregular area on the image resulting from detector saturation) obscured sample structure. In addition, the structure of the cells was altered after several such scans (this was determined by viewing the
Fig. 5. SE (a), CL (b), and overlay (c) images of a lyophilized powder of GFP (green fluorescent protein). Beam accelerating voltage = 10 keV, Z focus = 19 mm, magnification = 3.6 K. Bar = 10 lm.
same sample following gold sputter coating where these damaged areas of the sample could be identified in relation to adjacent undamaged cells which had not been irradiated). We tested fluorescent/nanogold dual-labeled probes (Fab’ fragments of polyclonal IgG covalently conjugated to a fluorescent dye molecule and a 1.4 nm gold particle) (Invitrogen, Carlsbad, CA) as a possible discharge mechanism with controlled, even and minimal distribution of the gold by necessarily binding to the antibody targets on the cell. This resulted in no reduction of charging but, in addition, we noted no loss of fluorescence such as reported by others in the use of gold coatings [2,3,19,20]. Therefore we tested osmium and gold coatings of various concentrations. Osmium provided little reduction of sample charging even to the degree that fluorescence was almost totally obscured. However, a 15 s sputter coating of gold–palladium (15 mA, 2.2 kV, using Argon plasma)
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(Bio-Rad, E-5400, Hercules, CA) greatly reduced the incidence of charging on the cells while passing about 50% of the emission observed without coating. A 60 s coating at the same power settings obliterated detectable fluorescence. Fo¨rster quenching distance calculations [20,21] show that a 12 nm or greater separation between a fluorophore and a gold particle prevents energy transfer/quenching. The continuity of gold particle deposition as well as the overall concentration and depth resulting from different power settings and sputter times should be considered in the determination of the most efficient level of gold deposition. Monocytes are a subset of mononuclear cells found in circulating blood. They are distinguished from other peripheral mononuclear cells (PBMNC) by the presence of a cell surface membrane-associated protein called CD14. Monocytes are also unique in that they possess extensive surface membrane which forms ‘‘veil-like’’ pro-
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trusions. Other PBMNC are more spherical and have short protrusions. A fluorescently tagged antibody to the CD14 protein, incubated with a sample of PBMNC, selectively labels the c cells and is routinely used in fluorescence activated cell scanning (FACS) to identify and select this population. CL detection could be used to grossly identify the CD14+ cells (Fig. 6b) as well as identify typical monocyte morphology and the distribution of surface proteins on the cells. Fig. 6 shows these morphological differences and the CL identification of the CD14+ cells. The fluorescence collected (b) from a select few of the cells in the field of Fig. 6a shows that the power of specific labeling, which is used routinely in confocal microscopy, is translatable to SEM-CL. As demonstrated in (c–f), regions bearing a high concentration of label can be associated with some SE differentiated structures such as protrusions (f, arrow) and veil edges.
Fig. 6. Cathodoluminescence images of fluorescent labels attached to human peripheral blood mononuclear cells. The cells were labeled with anti-human CD14-FITC/Alexa 656 and lightly sputter coated with gold after critical point drying. SE images (a, c), CL images (b, d). Images (a and b) are a field of cells of which only four have the veil-like shape and approximate size of CD14+ cells and display cathodoluminescence. Images (c and d) are magnifications of a similarly cathodoluminescent cell in this sample. Image (e) is an overlay of (d) on (c) and (f) is an enlargement of the boxed area in (e). Higher concentrations of the label occur on the edges of the veils and on some specific structures (arrow). Beam accelerating voltage = 5 keV, (a, b) Bar = 10 lm, (c–e) Bar = 600 nm, (f) Bar = 200 nm.
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5. Conclusion No effective method has previously been demonstrated to link single, or a combination of potentially interacting, molecules to SEM resolved cellular structures. We have combined CL with SE detection to image specific antibody labels on biological cells at the submicron scale. This will be particularly useful in visualizing tagged molecules on specific cell structures, such as veils, dendrites, or gap junctions, where a lowered concentration, or the absence, of a population of receptors or ligands might evince a pathological condition. During the optimization experiments we found that the best CL images were obtained with the highest current range of the electron probe, low beam acceleration voltages (3–5 keV) and a beam current near 10 lA. In addition, we found it necessary for samples to be well grounded to avoid charge buildup. Plating cells with a thin adherent material on an aluminum surface that can be firmly attached to the instrument stage aids in this precaution. An overcoat of gold of limited concentration can help in charge dissipation, but this must be monitored against a loss of fluorescence due to quenching. Fixatives must be thoroughly washed out before labeling to avoid chemical interactions and glutaraldehyde should be avoided due to its contribution to autofluorescence. The effectiveness of SEM-CL as a tool for studying biological samples can be improved. We must explore the relationship between accelerating voltage and depth of electron penetration and the effect of electron scattering on the size of the excitation volume. The emissive volume and its relationship to secondary or tertiary energy sources must also be measured. These aspects will more accurately locate the three-dimensional origin of a photonic signal. In addition, simultaneous detection of labels of multiple colors is a necessity in studies of biological interactions. An enhanced SEM-CL optimized for biological applications would incorporate a PMT of the highest sensitivity and widest spectral range available such as the Hamamatsu R928P which has, a single emission gathering umbrella sensitive to all the emission wavelengths and covering a maximum of the sample surface, simultaneous detection of X-rays, back-scattered electrons, secondary electrons and photons, and separation of the photonic signal by color. Such instrumentation, in combination with recently developed highly photostabile fluorophores and quantum dots would increase the opportunity of identifying molecular components of nanoscale morphological structures on
cells and tissues and their cellular distribution relative to each other. Acknowledgement This investigation was supported, in part, by Grants NCI P30 CA 15083, NIH P50 CA 108961, NCI P50 CA 91956-6 and by Mayo Foundation. We also thank Dr. Angela Klaus, Director of the Microscopy and Imaging Facility, American Museum of Natural History, New York, for support and encouragement. References [1] C.E. Norman, Microscopy and Analysis 53 (2002) 13. [2] E.F. Bond, D. Beresford, G.H. Haggis, Journal of Microscopy 100 (1974) 271. [3] W.A. Barnett, M.L. Wise, E.C. Jones, Journal of Microscopy 105 (1975) 299. [4] P.V.C. Hough, W.R. McKinney, M.C. Ledbetter, et al., Proceedings of the National Academy of Sciences of the United States of America 73 (1976) 317. [5] M.D. Muir, P.R. Grant (Eds.), Cathodoluminescence, Academic Press, London, 1974. [6] W. Brocker, G. Hauck, R. Blaschke, et al., Microscopica Acta Supplement 2 (1978) 260. [7] W. Brocker, G. Pfefferkorn, Scanning Electron Microscopy 2 (1979) 125. [8] A.S. Loginov, S.M. Chebanov, G.V. Saparin, et al., Scanning 20 (1998) 442. [9] G. Ning, T. Fujimoto, H. Koike, et al., Journal of Histochemistry and Cytochemistry 41 (1993) 617. [10] E. Kimura, T. Sekiguchi, H. Oikawa, Archives of Histology and Cytology 67 (2004) 263. [11] M. Cole, D. Ryer, Electro Optical Systems Design, Milton S. Kiver Publications Inc., 1972, pp. 16. [12] W.C. Chan, S. Nie, Science 281 (1998) 2016. [13] N. Panchuk–Voloshina, R.P. Haugland, J. Bishop–Stewart, et al., Journal of Histochemistry and Cytochemistry 47 (1999) 1179. [14] F.G. Prendergast, K.G. Mann, Biochemistry 17 (1978) 3448. [15] A. Einstein, Annual Physik 17 (1905) 132. [16] B. Hafner, Scanning Electron Microscopy Primer, University of Minnesota, Characterization Facility (2007). Available from: