Environmental scanning electron microscopy for biology and polymer science

environmental scanning electron microscopy Environmental scanning electron microscopy for biology and polymer science Debbie J Stokes, FEI Company, E...
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environmental scanning electron microscopy

Environmental scanning electron microscopy for biology and polymer science Debbie J Stokes, FEI Company, Eindhoven, The Netherlands Introduction Scanning electron microscopy (SEM) has become a routine tool for studying the micro- and nanostructure, composition and properties of bulk materials, with a large depth of field and a wide variety of signals giving the ability to visualise fine surface morphology and provide quantitative compositional information over comparatively large areas. SEM traditionally requires high vacuum conditions throughout the column, to minimise primary electron scattering and maintain a focused beam. Volatile components should not normally be present in a specimen for conventional SEM, as this would compromise the vacuum and put the electron source at risk of contamination. Many biological specimens, foams, emulsions, food systems and so on contain gases, water or oils and so require preparation to remove outgassing substances using techniques such as chemical fixing, alcohol dehydration, freeze-drying and critical point drying. Electron microscopy of specimens containing a substantial amount of liquid is usually performed using cryogenic procedures and cryo-SEM apparatus to render the specimen solid before and during imaging. These techniques can change the structural or chemical nature of the specimen, leading to potential artefacts, especially if not carried out with care. Electron irradiation can result in a build-up of negative charge unless the specimen is electrically conductive, in which case charge can be dissipated via a grounded specimen holder. Metallic samples can be imaged with ease in the SEM, but less conductive samples dissipate negative charge much less efficiently and therefore charge builds up, leading to deterioration in image quality. Insulating samples can be sputter-coated with a conduc-

tive material although, again, artefacts may be introduced and fine structural details obscured under the coating. Alternatively, a low energy electron beam (arbitrarily a few tens to a couple of thousand electronvolts) can be used in order to maintain a balance between primary and emitted electrons, eliminating the need for coating. However, it can be difficult to find the right criteria for charge balance when the specimen consists of regions with differing electrical properties. Ultimately, the desire to go beyond extensively prepared, static specimens, especially for biological and polymeric materials, became the driving force for the development of a new type of SEM in which gas is used to mitigate charging and facilitate alternative mechanisms for signal detection, allowing electrically insulating materials to be viewed without conductive treatment and also affording direct observation of moist or liquid specimens and dynamic phenomena. Terms that are commonly used are ‘variable pressure’, ‘lowvacuum’ and ‘environmental’ SEM, and these tend to be used in different contexts: variable pressure and low vacuum suggesting use of a gas for charge control (henceforth referred to as lowvacuum mode), environmental suggesting that there is an additional need for some specific gas (such as water vapour) at a certain pressure and/ or temperature (henceforth referred to as ESEM mode). The distinction between the two is usually clear from the context: polymeric materials are typically imaged under low-vacuum conditions, while many biological, moist or liquid specimens require a suitable water vapour environment to maintain thermodynamic equilibrium. Figure 1 helps to demonstrate how low-vacuum mode can be used to image an insulating sample that is otherwise very problematic in high vacuum.

Historical background From about the 1950s, differentially pumped, aperture-limited ‘environmental chambers’ were being tried for the TEM, as well as sealed containers with thin, electron-transparent windows (see, for example [1]). In 1970, Swift and Brown successfully demonstrated a version of the latter for the SEM in STEM mode, with the specimen contained under atmospheric pressure [2], and Lane described an aperture-limited ‘environmental control stage’ for the SEM, along with scattering cross-sections and mean free paths of electrons in various gases such as hydrogen, oxygen, nitrogen and the noble gases [3]. Robinson, Moncrieff and others in the 1970s adapted the SEM specimen chamber itself to accommodate imaging in gas, with the goal of imaging biological specimens without preparation [4]. Stable imaging of water was demonstrated by Robinson [5] using a modified JEOL JSM 2 SEM, containing a single pressure limiting aperture to separate the vacuum at the electron source from the specimen chamber accommodating pressures of up to 665 Pa (5 torr). It was observed that imaging uncoated insulators at pressures above about 10 Pa reduces the effects of specimen charging. Moncreiff et al. proposed that collisions between electronic species and gas molecules, resulting in the production of positive ions, provided the mechanism for the observed charge ‘neutralization’ [6], calculating the effects and amount of scattering of primary electrons in nitrogen gas [7]. An important conclusion of this work was that, although some primary electrons may be scattered tens to thousands of microns away from their original trajectories, the remaining electrons still form a focused probe for imaging. The scattered electrons reduce the total current in the focused probe Figure 1 Environmental scanning electron microscope images of an optoelectronic transistor, demonstrating low vacuum mode using water vapour at a pressure of 150 Pa (1.1 torr) for imaging an extremely electrically insulating specimen consisting of several polymeric layers sandwiched between two thin regions of evaporated gold on a glass substrate. By varying the primary beam energy (left = 15 keV, right = 30 keV), it is possible to selectively image either (left) the uppermost gate electrode or (right) the underlying source-drain (each 50 nm thickness) buried beneath an 800 nm thickness of polymer multilayers. In high vacuum mode, this uncoated specimen is difficult to image at low energy and impossible at higher energy. Horizontal field width = 3 mm. Sample courtesy of Catherine Ramsdale, Plastic Logic, Cambridge, UK.

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environmental scanning electron microscopy

while adding a uniform component (probe ‘skirt’) to the overall background signal. In 1978, Robinson commercialised this new technology via ETP Semra Pty Ltd, with a device later called the charge-free anti-contamination system (CFAS). They were mostly sold in Japan, attached to Akashi/ISI SEMs, and had a pressure limit of 270 Pa (2 torr). In 1980, Akashi/ISI integrated the CFAS and launched the WET SEM. Meanwhile, a two-aperture system giving three differentially pumped zones was introduced in 1979 by Danilatos and Robinson [8]. Shah and Beckett were obtaining similar results using their moist environment ambient-temperature SEM (MEATSEM) [9], and Neal and Mills built such a system using a Cambridge Stereoscan Mk II [10]. Danilatos, working with Electroscan Corp, developed the environmental SEM (ESEM) in the 1980s, with a pressure limit of 2.7 kPa (20 torr) and a secondary electron (SE) detector utilising the gas ionisation cascade to amplify SE signals [11]. By the 1990s, the list of commercial instruments included low vacuum SEM, (LVSEM, JEOL); natural SEM (NSEM, Hitachi); environment controlled SEM (ECO SEM, Amray Inc); and the EnVac (Gresham Camscan). In 1995 Leica and Carl Zeiss formed LEO Electron Microscopy Ltd, and introduced the variable pressure SEM (VPSEM), later adding extended pressure (EP) mode. VPSEM was then incorporated into Carl Zeiss SMT in 2001. In 1996, the controlled pressure SEM (CPSEM) was introduced by Philips Electron Optics. During the same year, Philips acquired Electroscan Corp and its ESEM technology group, and subsequently merged with FEI, becoming known as FEI Company in 2002. These commercial technology developments paved the way for researchers to expand both the applications and knowledge of the underlying principles of the technique. Results began to proliferate from about the late 1990s, with the most active research centred on the ESEM instrument in particular. Gas cascade signal amplification was investigated in more detail [12, 13] along with the properties of different imaging gases [14, 15], as well as spurious X-ray production emanating from primary electrons scattered by the gas [1621]. Some novel contrast mechanisms in solids, dependent on specific ESEM operating conditions and dielectric specimen properties, were increasingly explored [22-27] (see Fig 2) and similarly for heterogeneous liquids [28] (see Fig 3). The influence of positive ions generated in the gas cascade process was elucidated, further helping to explain some of the charge-related phenomena previously seen, as well as other effects [29-31], and evaluations and methodologies were developed for ESEM observation of liquid [32], polymeric [33, 34] and hydrated biological materials [35-39]. Imaging of biological and polymeric materials Modern instruments of this genre are capable of high resolution imaging and analysis in both vacuum and gas, becoming a widely adopted addition to the characterisation toolkit in facilities worldwide. An important feature is the capability to provide an appropriate chemical environment for a given specimen or experiment when required. For biological or hydrated materials, this typically involves using water vapour as the imaging gas along with specimen cooling, to reach the relevant thermodynamic conditions 68

Figure 2 ESEM micrograph of embedded, flat-polished grains of the mineral gibbsite (aluminium hydroxide) showing a region of enhanced contrast as a result of imaging conditions and dielectric properties of the specimen. Impurityrelated growth zones are uniquely revealed in this mode. Note how the application of a carbon coating (left) completely suppresses this form of contrast. Water vapour pressure = 150 Pa (1.1 torr). Scale bar = 20 µm. Courtesy of B. J Griffin, University of Western Australia.

Figure 3 Dynamic charging effects in a liquid-state oil-in-water emulsion. In the left-hand image, oil droplets appear dark in a continuous aqueous phase. When the scan rate is slowed by a factor of 60, oil droplets appear bright. This contrast reversal can be switched back and forth as the scan rate is varied, indicating differences in the dielectric properties of the liquids involved. Horizontal field width = 200 µm. Reproduced with permission from Ref [28]. Copyright John Wiley and Sons, Inc. Figure 4 Phase diagram for water showing both the 100% relative humidity (RH) curve (grey) and a modified curve (black) for an example specimen containing solutes, in thermodynamic equilibrium with its vapour at 75% RH, to demonstrate some applicable ranges of pressures and temperatures in ESEM mode. Points that lie above and below the curves represent condensing and evaporating conditions, respectively.

and so minimise or avoid unwanted drying of the specimen (in addition to ensuring that the specimen does not change appreciably during pumpdown from atmospheric conditions). In this way, ESEM imaging can be applied to, for example, cells/tissues and emulsions/suspensions, with little or no specimen preparation, or used to follow dynamic processes, with the caveat that delicate uncoated materials are more susceptible to radiation damage, which is exacerbated in the presence of reactive species, particularly those arising from electron irradiation of water [40, 41]. Initially, for the setting up of ESEM conditions,

the general rule-of-thumb was to follow the saturated vapour pressure curve for water and aim for thermodynamic parameters that enable adequate imaging. The imaging parameters are somewhat interdependent and largely based on the electron scattering and signal amplification properties of the gas, voltage on the detector anode, gas path length (final PLA-to-sample distance), sampleto-detector distance and electron emission coefficients of the specimen [13, 42]. With an on-axis detector, such as the environmental secondary detector (ESD) and its successor the gaseous secondary electron detector (GSED), the final PLA

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environmental scanning electron microscopy

forms part of the detector assembly, and so the gas path length (GPL) is roughly the same as the sample-to-detector distance. With this arrangement and water vapour as the imaging gas, typical pressures used in ESEM mode are in the region of 600-800 Pa (4.5-6 torr), combined with specimen cooling to temperatures around 0-4°C in order to maintain 100% relative humidity (RH) in accordance with the phase diagram for water (see Figure 4). It was increasingly appreciated that consideration of the specimen’s inherent thermodynamic equilibrium is needed, along with kinetics, and these lead to environmental conditions that lie below the curve for pure water, permitting either a lower pressure or higher temperature to be used. Working at lower pressures helps to reduce primary electron scattering, improving the signalto-background ratio and focused beam current for more effective imaging and x-ray microanalysis. Figure 4 shows a modified curve taking into account the equilibrium pressures and temperatures for a saturated solution containing NaCl, having equilibrium RH = 75%. Thus, for real biological and hydrated/liquid materials, which typically contain solutes such as salt, this allows for adjustments to operating parameters to improve performance without compromising specimen stability. Furthermore, working at pressures above the specimen’s equilibrium RH (i.e. simply following the 100% RH rule-of-thumb) can cause unwanted condensation of water onto the specimen [43, 44]. Hence, for a given specimen temperature, say 3°C, water vapour pressures can be lowered by some 300 Pa (roughly 2 torr) and, as water loss is kinetically slow at low temperatures, it may be possible to go to even lower pressures for short periods (minutes) without causing undue evaporation of water from the specimen. For a more detailed discussion, refer to [45]. Later refinements include placing the SE detector anode off-axis, with the option of fitting a PLA extending from the pole-piece towards the specimen, thus decoupling the specimen-detector signal amplification distance from the (decreased) gas path length of primary electrons, allowing greater flexibility and optimisation of operating parameters. This led to the introduction of the off-axis large field detector (LFD). In conjunction with a large aperture in the PLA, a higher field-of-view can be scanned, albeit with a lower maximum pressure limit. However, because the specimen-to-detector distance is independently increased, any reduction in SE signal caused by the lower gas concentration is compensated for by the longer path length in which gas cascade amplification can occur. For specimens such as non-hydrated materials, which generally only require pressures in the range of about 70-400 Pa (approx. 0.5-3 torr), the combination of LFD and low vacuum mode covers many use cases and is well suited to imaging of polymeric materials, including experiments of a dynamic nature. Examples of the latter include tensile testing of biopolymers [46, 47] and nanoparticles [48]. Other gases may also be used for imaging in low-vacuum mode, e.g. nitrogen, in the case that the specimen is sensitive to processes such as oxidation. At the other end of the spectrum is high pressure imaging (up to 2.7 kPa, 20 torr for water vapour) using the GSED, for SE detection, or the more recently introduced ESEM gaseous

Figure 5 Gaseous secondary electron detector (GSED) image of leather under conditions of room temperature and a water vapour pressure of 2 kPa (15 torr), giving RH approx. 65%, demonstrating very good resolution despite the high pressure/humidity. Courtesy of K. Mam, FEI Nanoport, Eindhoven, The Netherlands.

Figure 6 ESEM-STEM micrograph of a di-block co-polymer consisting of annealed polystyrene-polyisoprene. The lamellae are around 100 nm in width and are visible without preferential contrast-enhancing staining of either phase. Courtesy of L. Staniewicz, University of Cambridge, UK.

analytical detector (ESEM GAD), for SE and BSE detection with optimised geometry for X-ray microanalysis. These detectors operate well for conditions in which high humidity is attained through high pressures of water vapour imaging gas and little or no specimen cooling, lifting some of the restrictions on operating parameters and bringing environmental conditions a step closer to reality, depending on the relevant criteria for the experiment [49]. Figure 5 shows an example of the GSED operating at high pressure for imaging an organic material under high humidity conditions at room temperature. Another interesting development is the incorporation of detectors for collecting bright- and dark-field (BF and DF, respectively) transmitted electron signals through thin specimens, socalled STEM-in-ESEM or wetSTEM, which offer further potential for ESEM characterisation of polymeric systems and biological ultrastructure. The relatively low primary beam energies used in (E)SEM, 30 keV and below, lend themselves particularly well to the observation of organic materials. Primary electron scattering as the beam passes through a material (the mechanism for BF/DF imaging) increases exponentially with decreasing energy, and so the resultant signal is sensitive to small compositional changes at low primary energies, making the contrast difference between low density phases more pronounced. One advantage of this is that contrast-enhancing preferential staining of organic phases may not be necessary, as is often the case when working with a high-voltage S/TEM instrument. Figure 6 shows a di-block co-polymer in which its

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characteristic lamellar microstructure can be seen in the absence of staining. It is argued that traditional staining of this and similar systems for high vacuum EM studies leads to artefacts caused by the swelling of the stained phase and concomitant compression of the unstained phase [50]. Another advantage of STEM mode is that the interaction volume of primary electrons involved in BF/DF signal formation as the beam passes through a thin specimen is smaller than that of a beam spreading out and coming to rest within a bulk specimen, from which SE and BSE signals emanate, hence in principle BF/DF imaging gives higher resolution than SE/BSE imaging. STEM-in-ESEM typically refers to studies that involve gas as a means of charge control (in other words, low-vacuum mode), rather than for ‘environmental’ reasons, and has found particular use in the study of polymer films. For organic photovoltaic devices and LEDs, the morphology of semiconducting polymer blends can be related to device performance, where exciton diffusion lengths at polymer-polymer interfaces are critical. Both low-vacuum SE imaging of cross-sections [51] and STEM-in-ESEM plus X-ray microanalysis of films [52] have demonstrated that the technique is very well suited to the study of phase separated polymer domains, where parameters such as accelerating voltage and imaging mode can be flexibly adjusted to reveal the relevant features of interest. WetSTEM involves the same use of water vapour pressures and specimen temperatures for thermodynamic control as for ESEM mode, and has the potential for increasing the avail69

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able information at the nanoscale for materials interacting with an aqueous phase. The specimen (liquid or solid) is deposited on a TEM grid in contact with a cooled surface such as a Peltier stage. WetSTEM is exemplified by studies of polymer colloids or nanoparticles in aqueous suspension [53] (see Figure 7), nanoscale wetting dynamics of materials [54, 55] and controlled dehydration of bacteria [56]. Conclusions Bringing together the various approaches briefly outlined here, Figure 8 schematically depicts the configurations and conditions that contribute to the successful characterisation of specimens for biology and polymer sciences in ESEM. Over several decades, ESEM has developed and grown into a commonplace tool for observing a wide range of materials without the need for extensive specimen preparation. As such, it lends itself particularly well to electrically insulating, organic materials such as those in biology and polymer science, both for studying static microstructure and in situ dynamic processes that occur during, for example, cooling, heating, wetting, dehydration and mechanical testing. Current and future developments will continue to lead the technique in the direction of research in, for example, physiological studies including nanoparticle interactions with cells, structureproperty relationships in complex polymer composites or organic semiconducting materials and in situ chemical reactions such as catalytic growth of nanotubes, to name just a few. References

1. Swift, J.A. and A.C. Brown, Environmental cell for examination of wet biological specimens at atmospheric pressure by transmission scanning electron microscopy. J. Physics E: Scientific Instruments, 1970. 3(11): 924. 2. Parsons, D., Radiation damage in biological materials, In Physical Aspects of Electron Microscopy and Microbeam Analysis, B. Siegel, Editor. 1975, New York. p. 259-265. 3. Lane, W.C. The environmental cold stage. In Proceedings of the Third Scanning Electron Microscopy Symposium. 1970. IIT Research Institute, Chicago, IL, 60616 4. Robinson, V.N.E., The elimination of charging artifacts in the scanning electron microscope. J. Physics E: Scientific Instruments, 1975. 8(8): 638-640. 5. Robinson, V.N.E. A wet stage modification to a scanning electron microscope. in Proc. 8th Int. Cong. Electron Microscopy. 1974. Canberra: Aust. Acad. Sci. 6. Moncrieff, D.A., V.N.E. Robinson, and L.B. Harris, Gas neutralisation of insulating surfaces in the SEM by gas ionisation. J. Phys. D: Appl. Phys., 1978. 11(17): 2315-2325. 7. Moncrieff, D.A., P.R. Barker, and V.N.E. Robinson, Electron-scattering by the gas in the scanning electron microscope. J. Physics D: Applied Physics 1979. 12(4): 481-488. 8. Danilatos, G.D. and V.N.E. Robinson, Principles of scanning electron microscopy at high specimen chamber pressures. Scanning, 1979. 2: 72-82. 9. Shah, J.S. and A. Beckett, A preliminary evaluation of moist environment ambient temperature scanning electron microscopy (MEATSEM). Micron, 1979. 10: p. 13-23. 10. Neal, R.J. and A. Mills, Dynamic hydration studies in an SEM. Scanning, 1980. 3(4): p. 292-300. 11. Danilatos, G.D., U.S. Patent No. 4,897,545: Electron detector for use in a gaseous environment, 1990, ElectroScan Corporation (Danvers, MA). 12. Meredith, P., A.M. Donald, and B. Thiel, Electron-gas interactions in the environmental scanning electron microscopes gaseous detector. Scanning, 1996. 18(7): 467-473. 13. Thiel, B.L., et al., An improved model for gaseous


Figure 7 Annular dark field ESEM micrograph of an aqueous-phase homogeneous acrylic latex during evaporation by heating using wetSTEM. On the right hand side is a monolayer, while multiple particle layers are to the left. The large circle visible in the background is a hole in the carbon layer of the TEM grid. Scale bar = 500 nm. Copyright Elsevier. Reproduced with permission from ref [53].

Amplification in the environmental SEM. J. Microscopy 1997. 187(Pt. 3): 143-157. 14. Fletcher, A., B. Thiel, and A. Donald, Amplification measurements of potential imaging gases in environmental SEM. J. Phys. D: Appl. Phys., 1997. 30: 2249-2257. 15. Stowe, S.J. and V.N.E. Robinson, The use of helium gas to reduce beam scattering in high vapour pressure scanning electron microscopy applications. Scanning, 1998. 20: 57-60. 16. Griffin, B.J. and C.E. Nockolds. Quantitative EDS analysis in the ESEM using a bremstrahlung intensitybased correction for primary electron beam variation and scatter. Microscopy and Microanalysis ‘96. 1996. 17. Bilde-Sorensen, J. and C.C. Appel. X-ray spectrometry in ESEM and LVSEM: corrections for beam skirt effects. in SCANDEM-97. 1997. 18. Doehne, E., A new correction method for highresolution energy-dispersive X-ray analyses in the environmental scanning electron microscope. Scanning, 1997. 19(2): 75-78. 19. Mansfield, J.F., X-ray microanalysis in the environmental SEM: A challenge or a contradiction? Mikrochimica Acta, 2000. 132(2-4): 137-143. 20. Gauvin, R., Some theoretical considerations on X-ray microanalysis in the environmental or variable

pressure scanning electron microscope. Scanning, 1999. 21(6): 388-393. 21. Newbury, D.E., X-ray microanalysis in the variable pressure (environmental) scanning electron microscope. Journal of the National Institute of Standards and Technology, 2002. 107(6): 567-603. 22. Clausen, C. and J. Bilde-Sorensen, Observation of voltage contrast at grain boundaries in YSZ. Micron and Microscopia Acta, 1992. 23(1/2): 157-158. 23. Horsewell, A. and C. Clausen. Voltage contrast of ceramics in the environmental SEM in ICEM 13. 1994. Paris, France. 24. Griffin, B.J., A new mechanism for the imaging of non-conductive materials: an application of chargeinduced contrast in the environmental scanning electron microscope. Microscopy & Microanalysis, 1997 3 (Suppl. 2): 1197-1198. 25. Doehne, E., Charge contrast: Some ESEM observations of a new/old phenomenon. Microscopy & Microanalysis, 1998. 4(Suppl.2: Proceedings): 292-293. 26. Baroni, T.C., et al., Correlation between charge contrast imaging and the distribution of some trace level impurities in Gibbsite. Microsc. Microanal., 2000. 6: 49-58. 27. Clode, P.L., Charge contrast imaging of biomaterials in a variable pressure scanning electron microscope. Journal of Structural Biology, 2006. 155(3): 505-511. 28. Stokes, D.J., B.L. Thiel, and A.M. Donald, Dynamic secondary electron contrast effects in liquid systems studied by environmental SEM (ESEM). Scanning, 2000. 22(6): 357-365. 29. Craven, J.P., et al., Consequences of positive ions upon imaging in low vacuum SEM. J. Microscopy, 2002. 205(1): 96-105. 30. Toth, M., et al., Electric fields produced by electron irradiation of insulators in a low vacuum environment. J. Applied Physics, 2002. 91(7): 4492-4499. 31. Toth, M., et al., Quantification of electron-ion recombination in an electron-beam-irradiated gas capacitor. J. Physics D: Applied Physics, 2002. 35(14): 1796-1804. 32. Stokes, D.J., B.L. Thiel, and A.M. Donald, Using secondary electron contrast for imaging water-oil emulsions in the environmental SEM (ESEM). Microscopy & Microanalysis, 1998. 4(Suppl. 2: Proceedings): 300-301. 33. Cameron, R.E., Environmental Scanning Electron Microscopy in Polymer Science. Trends in Polymer Science, 1994. 2(4): 116-119. 34. Meredith, P. and A.M. Donald, Study of ‘wet’ polymer latex systems in environmental scanning electron microscopy: some imaging considerations. J. Microscopy, 1996. 181(1): 23-35. 35. Cameron, R.E. and A.M. Donald, Minimising sample evaporation in the environmental scanning electron microscope. J. Microscopy 1994. 173(3): 227-237.

Figure 8 Simplified schematic diagram (not to scale) showing a number of configurations for ESEM and low-vacuum imaging. On the left, with an on-axis detector and final pressure limiting aperture (PLA) close to the lens, the gas path length (GPL) and specimen-detector distance are roughly the same. The working distance can be varied to improve some of the factors involved in imaging, but at the expense of others. On the right, these parameters are decoupled, giving greater flexibility in operating parameters. For example, the GPL can be kept short by optionally extending the PLA towards the specimen, reducing the scattering of primary electrons, while the detector can be placed in an on- or off-axis position with various geometries to maximize the gas amplified signal. The geometry on the right is also optimised for a fixed X-ray take-off angle (e.g. 10 mm working distance). 25th Anniversary Issue September 2012 | MicroscopyandAnalysis

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36. Farley, A.N., A. Beckett, and J.S. Shah, MEATSEM (Moist Environment Ambient-Temperature Scanning Electron-Microscopy) or the SEM of plant materials without water-loss. Institute of Physics Conference Series, 1988(93): 111-112. 37. Farley, A.N., A. Beckett, and J.S. Shah. Comparison of beam damage of hydrated biological specimens in high-pressure scanning electron microscopy and lowtemperature scanning electron microscopy. In Proc. XIIth International Congress for Electron Microscopy. 1990, San Francisco Press. 38. Gilpin, C. and D.C. Sigee, X-ray microanalysis of wet biological specimens in the environmental scanning electron-microscope .1. Reduction of specimen distance under different atmospheric conditions. Journal of Microscopy 1995. 179: 22-28. 39. Stokes, D.J., Characterisation of soft condensed matter & delicate specimens using environmental scanning electron microscopy (ESEM). Advanced Engineering Materials, 2001. 3(3): 126-130. 40. Kitching, S. and A.M. Donald, Beam damage of polypropylene in the environmental scanning electron microscope: an FTIR Study. J. Microscopy 1998. 190: 357-365. 41. Royall, C.P., B.L. Thiel, and A.M. Donald, Radiation damage of water in environmental scanning electron microscopy. J. Microscopy, 2001. 204(3): 185-195. 42. Thiel, B.L., Master curves for gas amplification in low vacuum and environmental scanning electron microscopy. Ultramicroscopy, 2004. 99(1): 35-47. 43. Tai, S.S.W. and X.M. Tang, Manipulating biological samples for environmental scanning electron microscopy observation. Scanning, 2001. 23: 267-272. 44. Muscariello, L., et al., A critical review of ESEM applications in the biological field. Journal of Cellular Physiology, 2005. 205: 328-334. 45. Stokes, D.J., Principles and practice of variable pressure/environmental scanning electron microscopy (VP-ESEM). 2008, Chichester: John Wiley & Sons. 46. Rizzieri, R., F.S. Baker, and A.M. Donald, A Study of

the large strain deformation and failure behaviour of mixed bioploymer gels via in situ ESEM. Polymer, 2003. 47. Rizzieri, R., et al., Superficial wrinkles in stretched, drying gelatin films. Langmuir, 2006. 22(8): 3622-3626. 48. Liu, T., A.M. Donald, and Z. Zhang, Novel manipulation in environmental scanning electron microscope for measuring mechanical properties of single nanoparticles. Materials Science and Technology, 2005. 21(3): 289-294. 49. Toth, M., et al., Secondary electron imaging at gas pressures in excess of 1k Pa. Applied Physics Letters, 2007. 91(Article No. 053122). 50. Staniewicz, L. et. al, The effect of osmium staining on lamellar spacing in thin polystyrene-polyisoprene diblock copolymer films. (Manuscript in preparation). 51. Ramsdale, C.M., et al., ESEM imaging of polyfluorene blend cross-sections for organic devices. Physica E: Low-dimensional Systems and Nanostructures, 2002. 14(1–2): 268-271. 52. Williams, S.J., et al., Imaging of semiconducting polymer blend systems using environmental scanning electron microscopy and environmental scanning transmission electron microscopy. Scanning, 2005. 27(4): 190-198. 53. Bogner, A., et al., Wet STEM: A new development in environmental SEM for imaging nano-objects included in a liquid phase. Ultramicroscopy, 2005. 104(3-4): 290-301. 54. Barkay, Z., Wettability study using transmitted electrons in environmental scanning electron microscope. Applied Physics Letters, 2010. 96(18): 183109-183109-3. 55. Rykaczewski, K. and J.H.J. Scott, Methodology for imaging nano-to-microscale water condensation dynamics on complex nanostructures. ACS Nano, 2011. 5(7): 5962-5968. 56. Staniewicz, L., et al., The application of STEM and in situ controlled dehydration to bacterial systems using ESEM. Scanning, 2012 (in press).


Debbie Stokes has a PhD in physics from the Cavendish Laboratory in Cambridge, UK. Debbie is the Honorary Secretary for Physical Science at the Royal Microscopical Society and author of the handbook ‘Principles and Practice of Variable Pressure/ Environmental Scanning Electron Microscopy (VP-ESEM)’. Recent projects include the Titan ETEM at FEI Company. abstract Electron microscopy of materials in a gaseous environment has its origins as far back as about the 1950s for the transmission electron microscope (TEM), later followed by developments for the scanning electron microscope (SEM). Focusing on the latter, this review outlines the history and commercial development of the technology, culminating in the environmental SEM (ESEM), and the techniques for applying ESEM to materials of interest in biology and polymer sciences. corresponding author details Dr Debbie J Stokes, FEI Company, Achtseweg Noord 5, 5651GG Eindhoven, The Netherlands Tel: +31 40 23 56000 Email: [email protected] Microscopy and Analysis 26(6):67-71 (AM), 2012

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