- Email: [email protected]
Environmental Scanning Electron Microscopy
2.21
Environmental Scanning Electron Microscopy
AM Donald, University of Cambridge, Cambridge, UK © 2012 Elsevier B.V. All rights reserved.
2.21.1 2.21.2 2.21.3 2.21.4 2.21.4.1 2.21.4.2 2.21.5 2.21.6 References
Introduction The Instrument: A Comparison with Conventional SEM Static Experiments Dynamic Experiments Hydration and Dehydration Mechanical Testing Biopolymers and Biofilms Conclusions
2.21.1 Introduction The technique of environmental scanning electron microscopy (ESEM) has been around for nearly 20 years. However, in many ways it is still in its infancy, and applications to polymer science remain relatively few, despite the advantages it may potentially confer. In part this reflects the fact that for so many purposes conventional scanning electron microscopy (CSEM) looks suf ficient and is familiar, widely available, and easy to use (see Table 1 and Reference 1). In part it would appear to have been due to failing marketing strategies within the companies that manufacture the different varieties of the instrument, initi ally implying the instrument was much simpler to use than many researchers found it, and subsequently failing to appreci ate fully its strengths for hydrated samples such as latex dispersions. However, it also has to be recognized that there are limitations of the instrument, as will be described later in this chapter, and some of these, including the fundamental challenge of beam damage and resolution when performing energy-dispersive X-ray (EDX) analysis, are likely always to limit its utility for certain types of sample and problem. Nevertheless, there are many advantages that can usefully be factored in for polymer scientists trying to study a wide range of different situations, and this chapter will seek to identify the special strengths of the technique.
2.21.2 The Instrument: A Comparison with Conventional SEM There are two key differences with regard to a conventional SEM (CSEM): the chamber around the sample does not need to be under high vacuum and, as a direct consequence of this, insulators do not need to be coated with a conductive coating. These key differences lie at the heart of all the potential advan tages of the instrument. The basic design is essentially the same as a CSEM: there is an electron source, which may (as in CSEM) be a simple tungsten filament, a LaB6 filament, or a field emis sion gun (FEG), each with its own advantages of brightness, durability, and cost. This gun operates under the standard conditions of high vacuum, although exactly how high a vacuum is required depends on the type of gun used. The electron beam travels down the column toward the sample chamber. However, in a CSEM this entire path, with lenses to Polymer Science: A Comprehensive Reference, Volume 2
539 539 541 542 542 543 543 544 544
focus the beam, is under the same high vacuum conditions as the gun, whereas this is not the case in the ESEM. Instead, the column has a series of different pressure zones, as shown in Figure 1, each separated from its neighbor by a pressure limit ing aperture (PLA) across which differential pumping operates. As a result, the pressure increases steadily down the column. This permits the last region, which is the sample chamber itself, to contain a significant pressure of gas, up to around 10–15 torr (∼ 2000 Pa). The upper limit for this pressure is set by a number of considerations including the desired resolution, the type of detector used, and the accelerating voltage. The presence of the gas degrades the image quality less than that which might be expected due to scattering of the incident electron beam by the gas molecules before the electrons ever reach the sample. In practice, as long as the working distance (the distance from the bottom of the last aperture to the sample) is kept small, and the pressure in the chamber is not too large, most electrons are not scattered and those that are, are scattered only a few times; this means beam broadening is not too much of a problem and the shape of the incident beam is still strongly peaked above a rather flat background, as shown in Figure 2. There will be some small signal arising from the further reaches of this flat background, but the majority of the signal will be initiated by the sharp peak at the center. The net effect is to introduce what is essentially a background DC signal on the signal of interest. Quoted instrumental resolution may be < 10 nm, but for polymers this is very unlikely to be realized because of the usual problems of electron beam damage. The type of experiments for which the existence of the broad back ground ‘skirt’ is likely to matter is when X-ray microanalysis is attempted.2 It is the presence of the gas around the sample which makes the key difference with CSEM, and which confers ESEM’s many advantages. Frequently – but not necessarily – this gas is water vapor. If it is, it in principle permits a hydrated sample to be maintained in its hydrated state, and this will be discussed further below. For polymers and other insulators, however, the presence of this ‘imaging gas’ confers a particular advantage because of the way it can reduce the usual problem of charging. In a CSEM, this is overcome by coating the surface of the sample, but the disadvantage of so doing is that fine surface features may be obscured. (Modern low-voltage instruments can to some extent overcome this problem of charging.) In the ESEM, coating is not usually required, although many
doi:10.1016/B978-0-444-53349-4.00044-3
539
540
Table 1
Structure Characterization in Real Space | Environmental Scanning Electron Microscopy
Comparison of conventional and environmental SEM CSEM
ESEM
Gun Sample environment
Tungsten, LaB6, or field emission High vacuum
SE Detector BSE detector X-ray detection Imaging of insulators
Everhart-Thornley Yes Yes Need conductive coating, unless working at very low voltages Only after dehydration
Tungsten, LaB6, or field emission Gas (typically water), pressures up to ∼ 2000 Pa with optional high vacuum Gaseous secondary electron detector Not yet very good at discriminating Yes, but rather poor spatial resolution Yes
Imaging of hydrated samples
Yes
Pressure 10–7 torr
Pressure zone Gun chamber
10–6 torr
Upper column
Anode
10–4 torr
EC2
Column
10–1 torr
EC1
10 torr
Specimen
Gun
Projection apertures Column valve Upper pressure limiting aperture Vacuum manifold
Specimen
Lower pressure limiting aperture and integrated detector
Figure 1 Schematic drawing of the ESEM column, showing the different pressure zones. 1 torr ∼ 133 Pa.
(a)
Primary beam
(b)
Figure 2 In the so-called oligo-scattering regime, electrons in the inci dent probe undergo few, if any, collisions. (a) Schematic of the scattering undergone by the incident beam (heavy arrowed line) in this regime. (b) The resultant probe has a sharp intensity peak superimposed on a low background ‘skirt’.
inexperienced users do not appreciate this fact and may still apply such a coating. But in practice the gas molecules in the sample chamber can be ionized by electrons emitted by the sample (secondary electrons (SEs) predominantly, since their
collision cross section with the gas molecules is higher than the more energetic backscattered electrons (BSEs)). As a conse quence of these ionizing collisions, there is a resultant cascade of electron–gas molecule ionizations which is shown schematically in Figure 3. Each ionizing collision produces a positively charged ion together with an additional so-called daughter electron, and this leads to amplification of the electrons traveling toward the detector. The magnitude of this amplification can be very significant.3 However, in the context of charging, what matters is the generation of a substantial number of positive gaseous ions. These tend to drift toward the negatively charged sample – negatively charged because of the implantation of incident electrons – and as they accumulate at the surface, they tend to compensate for the negative charge buildup, and so reduce the effect of charging. The processes involved are complex, and the interested reader should look at the review4 for more details. In general, however, the effect means that insulators do not need a conductive coating. Furthermore, if a new surface is opened up, for instance during a dynamic experiment to study fracture, charging is still not an issue; examples of this will be given below. In general, this means that ESEM is particularly powerful for dynamic
Structure Characterization in Real Space | Environmental Scanning Electron Microscopy
541
Positively charged detector Electron beam Cascade electrons
Vo
A
Positivion
+ Gaseous atom V
Earthed sample Figure 3 Schematic representation of the cascade amplification process that operates when a gas is present in the sample chamber. Electrons emitted from the sample collide with gaseous atoms and can ionize them, producing additional daughter electrons that contribute to the signal detected at the positively charged detector.
experiments of insulators. In addition, the absence of a coating means that the intrinsic contrast from the sample is not obscured as the electrons emerge from the sample surface: for polymers, with their typical low atomic numbers and small differences between chemically distinct species, this can confer a further advantage that has been little exploited in the literature. So there are two fundamental advantages of the ESEM over CSEM: the sample can be imaged in a hydrated state, and insulators do not need to be coated. Let us now look at the signal that is detected. In CSEM, the detector works by having a positive bias to which the electrons are attracted. Conventionally, there are two detectors for detecting the low-energy SEs, usually defined as those with energies < 50 eV, and the high-energy BSEs. The latter are electrons deflected through close to 180° on hitting the sample, whereas the former are electrons emitted from the sample due to inelas tic processes (ionization, etc.). Because the SEs possess low energy, they can only escape from close to the sample surface, and, therefore, are particularly useful at providing topographic information. In contrast, BSEs can come from much deeper within the sample, and the number generated depends on atomic number. Thus BSEs typically can provide information on chemical composition. In CSEM, the two detectors are set up to distinguish the different signals, and, therefore, topo graphic and chemical information can be obtained separately. However, these standard detectors do not work in the low vacuum conditions of an ESEM, and existing detectors for the ESEM are much less successful at distinguishing the different signals. An additional problem arises because of the nature of
the cascade amplification itself. This process amplifies all signals, thus amplifying noise such as that generated by BSEs hitting other parts of the chamber and causing additional electrons to be generated which contain no useful information. These electrons are known as SE2s, and tend simply to degrade image quality. A discussion of signal composition can be found in Reference 5. In practice, good BSE detectors are still not available for the ESEM, and the SE detector will not completely discriminate between topographic and compositional contrast. Finally, the ever-present problem of electron beam damage should be discussed. The origins of beam damage for polymers in conventional microscopy have been amply discussed in previous articles.6,7 For ESEM, there are some additional issues that need to be borne in mind, notably that there can be additional damage mechanisms, particularly in the presence of water. Relatively little work has been done specifically study ing these effects,8 but the work that has been done has demonstrated that beam damage is worse because of the free radicals generated by collisions between the water molecules and the electrons,9,10 and these free radicals can diffuse across the sample surface causing additional damage to that directly caused by the incident electrons.9,10
2.21.3 Static Experiments As indicated above, the presence of the gas in the chamber means that typically charging is no longer an issue for insula tors, as in CSEM, so that these samples can be studied without a conducting coating obscuring surface detail. Nevertheless, there
542
Structure Characterization in Real Space | Environmental Scanning Electron Microscopy
is no doubt that the presence of a thin gold layer may ‘sharpen’ the image, and it is presumably for this reason that some papers still adopt this approach.11,12 However, for other applications, the absence of a coating may reveal contrast otherwise obscured by it. This was initially observed for a nonpolymeric sample of water–oil emulsions.13 The origin of the contrast lies in the differences in electronic properties between the two components, and the consequent mechanisms by which electrons traveling to the surface can lose energy. Similar con trast effects have also been observed in ceramics such as gibbsite,14 although with a subtly different interpretation. For polymers, this has particular advantages because differences in atomic number are usually tiny, although some attempts have been made to explore chemical differences with BSEs.15 For instance, in block copolymers of polystyrene-polyisoprene, only carbon and hydrogen are pre sent just arranged in different ways and with slightly different densities. Nevertheless, clear contrast can be seen between the phases (Figure 4) without the customary use of staining to reveal the microstructure, for instance via the use of osmium tetroxide. Indeed, recent work is indicating that the use of such stains may actually preferentially swell one of the phases, meaning that measurement of lamellae spacings is inaccu rate,16 thus highlighting the particular value the minimal sample preparation required for imaging in the ESEM can bring. However, since the source of the contrast is related to the energy loss processes that occur as the electrons travel from their site of generation to the sample surface, the thickness of the material (for a thin film) through which the electrons must
2 µm
Figure 4 Contrast is visible without staining in a film of polystyrene-polyisoprene block copolymer with 50% of each component present. The polystyrene lamellae appear darker than the polyisoprene due to density differences. Micrograph courtesy of AJ Ryan and C Salou (University of Sheffield).
travel will play a part. So where there is significant topography present, care must be taken in order to interpret the images correctly. This has been demonstrated for the case of semicon ducting polymer blends spin cast from solution, in which significant topography is introduced during the drying process. Strong contrast is seen, but thought must be given as to which phase is identified with which polymer.17
2.21.4 Dynamic Experiments 2.21.4.1
Hydration and Dehydration
The ability to maintain a sample in a hydrated state – or indeed saturated with some fluid other than water – has value for many polymer applications, including those relevant to filtra tion and textiles. These appear to be the fields – other than biopolymer applications discussed further below – which have received most attention, but limited work has been carried out where a conventional polymer is explored in a hydrated state.18 The ability to change the level of hydration – either by deposit ing water on a sample or by dehydrating the sample and following the consequent morphological changes – opens up even more avenues. This can be done by changing the water vapor pressure in the chamber. Figure 5 shows the variation of the saturated vapor pressure (SVP) for water as a function of temperature, and indicates how small variations in temperature can shift the sample from hydrating to dehydrating conditions. Typically, the work needs to be done below room temperature, otherwise the requisite pressures are too high for useful ima ging, as can be inferred from Figure 5, but this does not alter the basic utility of this approach. For textiles, this has led to studies of swelling19–22 and analysis of the shape of water droplets21 and contact angle measurement of the droplets on the fibers.23,24 Although it might be thought that optical ima ging provided quite enough resolution to enable the contact angle to be measured accurately on a textile fiber, in practice Fresnel fringes often obscure the actual water–fiber interface, rendering precise measurement difficult, plus in principle local heterogeneities – leading to local variations in contact angle – can be explored, although this route does not seem to have been exploited. For membranes, some work has fully exploited the capabil ities of the ESEM to carry out dynamic experiments. For instance, as with the fiber work described above, Yu et al.25 have used the ability both to condense water droplets onto membranes to characterize their hydrophilicity/hydrophobi city and to explore the resulting contact angles. Wang et al.26 controlled the chamber relative humidity to permit hydration/ dehydration experiments to be carried out while observing the porosity of the membranes. Le-Clech et al.27 compared differ ent microscopic approaches for the study of biofouling of membranes, a very important problem in many applications, and noted that, although ESEM provided a major advance in being able to image the samples in appropriate wet mode, it was hard to focus on the deposited (alginate) films to observe fine structure. This will always be a challenge for ESEM, so it is a technique that most usefully works in tandem with other approaches in a complementary way. Cryogel structure during drying was also studied for polyacrylamide gels with different degrees of cross-linking so that the pore structure could be clearly visualized.28
Structure Characterization in Real Space | Environmental Scanning Electron Microscopy
543
25
Pressure (Torr)
20
Liquid
15 Condense
Evaporate
10 5 0
Gas
0
5
10
15
20
25
Temperature (°C) Figure 5 The SVP curve for water vapor, indicating the regimes in which evaporation or condensation will occur.
Additionally, solvents other than pure water can be used. This was done by Zhong et al. looking at aggregation of a terpolymer based on acrylamide in solutions of pure water and with salt present29 and with colloidal PMMA,30 in which the effect of the modification of the electrostatic screening due to the presence of salt was studied. This latter study highlights one particular class of polymeric samples, that of latices, that have been extensively studied during dehydration in the ESEM to reveal the stages in morphological development including the kind that make up paint formulations. The first researcher to carry out such experiments was Eckersley,31 and since then a range of different formulations have been explored, and novel insight obtained on the film drying process.32–37 In particular, a combination of ESEM with ellipsometry allowed the identifica tion of an intermediate stage in the standard stages of the drying process, in which most of the interstitial water had been lost, but complete densification was not achieved.38 Another variant of ESEM has also been used to study latex formulations by Bogner’s group. In this case, a wet-STEM (scan ning transmission electron microscopy) detector was used, so that the image was formed below the sample and the aggrega tion was observed.39
2.21.4.2
Mechanical Testing
As mentioned above, the relaxation of the condition for coating a sample surface means that ESEM offers particular attractions for performing mechanical tests in situ. Despite this, many of the papers that utilize ESEM in the context of mechanical testing only use the technique to study postmortem fracture surfaces. Rather few groups working on polymers have estab lished the requisite testing rigs. In general, these will need a cooling module as well, for the reasons given above about the useful SVP curves for water, but also because for polymers with glass transition temperatures close to room temperature, accu rate temperature control is essential if meaningful results are to be obtained.40 Different testing geometries can also be con structed: for instance Nase et al. have demonstrated the ability to carry out in situ peel tests on polymer bilayers,41 and Dragnevski et al. have produced a rig that allows for large extensions of up to more than 100% to be achieved.42
But the bulk of the work in the literature has used simple tensile/compression tests, allowing correlation of the deforma tion with the load-extension curves.43–48 In this way, it is possible to identify the nature of the failure route, or the loca tion of first yield in a heterogeneous sample. It would seem there is ample scope to extend the fairly limited studies that have been carried out to date.
2.21.5 Biopolymers and Biofilms The examples chosen so far have been focused on synthetic polymers of different types. In fact, ESEM has been particularly widely applied to biopolymers and biologically derived sam ples including biofilms, tissues, and bacteria, all of which contain substantial polymeric components. Cellulose has been explored in various states, both native fibers and wood and also dispersions of cellulose microfibrils. For all these types, the ability ‘not’ to dehydrate the sample has proved beneficial. In the case of microfibrillar assembly, how ever, it has also been the case that studying the arrangements of the microfibrils in dispersion49 and the way they aggregate has been possible at quite high resolution even in water. Many natural fibers have been studied ranging from hemp,50,51 through flax,52 including in situ mechanical testing,53 to wood. For the latter, there have been extensive studies on optimization of imaging54 and in situ swelling,55 and most notably, micromechanical studies.56–59 That the wood does not need to be dried or coated has clearly been identified as a key advantage for these studies. Composites containing cellu lose fibers have also been studied via ESEM, notably involving bacterial (rather than plant) produced cellulose.60,61 Food – broadly defined – is another area in which the ability to study wet systems has proved very useful. A recent review by James62 covers work in the field. Food here covers both vegetables (which could also be included in the cellulose section above) and starch-based materials, but also systems containing proteins and fats, although the last two categories have been much less exten sively studied than the former. The study of vegetables in the ESEM goes back to Uwins in 1993,63 who compared different sample preparation routes for potatoes, including the minimal preparation required for ESEM. In situ deformation of carrots
544
Structure Characterization in Real Space | Environmental Scanning Electron Microscopy
under different geometries64,65 and, more recently, onions66,67 has been used to look at failure modes correlated with load-extension data. Starch-based and starch-containing foods have been studied with various different motivations ranging from simply cataloging samples that had not been extensively treated prior to observation,68–70 to following hydration in situ by changing the sample chamber pressure,71,72 to carrying out in situ fracture tests.73 Rather less work has been done on other types of systems, although both fat-containing74,75 and fat-replacement systems43 have been looked at.
2.21.6 Conclusions ESEM offers many advantages for the polymer scientists, most notably the ability to image samples without applying a con ductive coating and the possibility of carrying out dynamic experiments – in situ hydration, dehydration, or mechanical tests. However, only certain parts of the community appear to have appreciated the potential of the technique fully, and there are many other types of sample upon which few ESEM experiments have been carried out. There are undoubtedly limitations, including beam damage (particularly in the pre sence of water vapor and the consequent generation of damaging free radicals) and, by not applying a coating, images may frequently look less sharp as if the resolution is worse than it truly is. Nevertheless, there are many advantages which the technique can confer, and it is to be hoped it will find an increasingly important place in the armory of characterization techniques used by the polymer science community.
References 1. Goldstein, J. I.; Newbury, D. E.; Joy, D. C.; et al. Scanning Electron Microscopy and X-ray Microanalysis, 3rd ed.; Springer: New York, 2002. 2. Mansfield, J. F. Mikrochim. Acta 2000, 132, 137–143. 3. Thiel, B.; Bache, I. C.; Fletcher, A. L.; et al. J. Micros. 1997, 187, 143–157. 4. Thiel, B. L.; Toth, M. J. Appl. Phys. 2005, 97, 051101. 5. Fletcher, A. L.; Thiel, B. L.; Donald, A. M. J. Micros. 1999, 196, 26–34. 6. Tsuji, M. In Comprehensive Polymer Science; Allen, G., Bevington, J. C., Eds.; Pergamon: Oxford, 1989; Vol. 1, p 803. 7. Sawyer, L. C.; Grubb, D. T.; Meyers, G. F. Polymer Microscopy, 3rd ed.; Springer: New York, 2008. 8. Zankel, A.; Chernev, B.; Brandl, C.; et al. Macromol. Symp. 2008, 265, 156–165. 9. Kitching, S.; Donald, A. M. J. Micros. 1998, 190, 357–365. 10. Royall, C. P.; Thiel, B. L.; Donald, A. M. J. Micros. 2001, 204, 185–195. 11. Sun, J.; Chen, X. S.; Lu, T. C.; et al. Langmuir 2008, 24, 10099–10106. 12. Guo, Z. P.; Li, Y. H.; Tian, H. Y.; et al. Langmuir 2009, 25, 9690–9696. 13. Stokes, D. J.; Thiel, B. L.; Donald, A. M. Langmuir 1998, 14, 4402–4408. 14. Watt, G. R.; Griffin, B. J.; Kinny, P. D. Am. Mineral. 2000, 85, 1784–1794. 15. Franz, N.; Ahlers, M. O.; Abdullah, A.; Hohenberg, H. J. Mater. Sci. 2006, 41, 4561–4567. 16. EMAG. J. Phys. Conf. Series. 2009, 241, (012077). 17. Williams, S. J.; Donald, A. M.; Thiel, B. L.; Morrison, D. E. Scanning 2005, 27, 190–198. 18. Miller, A. F.; Cooper, S. J. Langmuir 2002, 18, 1310–1317. 19. Jenkins, L. M.; Donald, A. M. Scanning 1997, 19, 92–97. 20. Jenkins, L. M.; Donald, A. M. Textile Res. J. 2000, 70, 269–276. 21. He, C. C.; Gu, Z. Y.; Tian, J. Y. J. Appl. Polym. Sci. 2003, 89, 3939–3943. 22. Wei, Q. F.; Wang, X. Q.; Mather, R. R.; Fotheringham, A. F. Fibres Text. East. Eur. 2004, 12, 79–83. 23. Jenkins, L. M.; Donald, A. M. Langmuir 1999, 15, 7829–7835. 24. Wei, Q. F.; Mather, R. R.; Fotheringham, A. F.; Yang, R. D. Textile Res. J. 2003, 73, 557–561.
25. Yu, H. M.; Schumacher, J. O.; Zobel, M.; Hebling, C. J. Power Sources 2005, 145, 216–222. 26. Wang, J.; Dismer, F.; Hubbuch, J.; Ulbricht, M. J. Membr. Sci. 2008, 320, 456–467. 27. Le-Clech, P.; Marselina, Y.; Ye, Y.; et al. J. Membr. Sci. 2007, 290, 36–45. 28. Plieva, F. M.; Karlsson, M.; Aguilar, M. R.; et al. Soft Matter 2005, 1, 303–309. 29. Zhong, C. R.; Ye, L.; Dai, H.; Huang, R. H. J. Appl. Polym. Sci. 2007, 103, 277–286. 30. He, C.; Donald, A. M. Langmuir 1996, 12, 6250–6256. 31. Eckersley, S. T.; Rudin, A. Prog. Org. Coat. 1994, 23, 387–402. 32. Keddie, J.; Meredith, P.; Jones, R.; Donald, A. In ACS Symposium Series; Provder, T., Winnik, M., Urban, M., Eds.; ACS: Washington, DC, 1996; Vol. 648, pp 332–348. 33. Meredith, P.; Donald, A. M. J. Micros. 1996, 181, 23–35. 34. Donald, A. M.; He, C.; Royall, C. P.; et al. Colloids Surf., A 2000, 174, 37–53. 35. Royall, C. P.; Donald, A. M. Scanning 2002, 24, 305–313. 36. Kugge, C.; Craig, V. S. J.; Daicic, J. Colloids Surf., A: Physicochem. Eng. Aspects 2004, 238, 1–11. 37. Dragnevski, K. I.; Donald, A. M. Colloids Surf., A: Physicochem. Eng. Aspects 2008, 317, 551–556. 38. Keddie, J. L.; Meredith, P.; Jones, R. A. L.; Donald, A. M. Macromolecules 1995, 28, 2673–2682. 39. Bogner, A.; Guimaraes, A.; Guimaraes, R. C. O.; et al. Colloid. Polym. Sci. 2008, 286, 1049–1059. 40. Zankel, A.; Poelt, P.; Gahleitner, M.; et al. Scanning 2007, 29, 261–269. 41. Nase, M.; Zankel, A.; Langer, B.; et al. Polymer 2008, 49, 5458–5466. 42. Dragnevski, K. I.; Fairhead, T. W.; Balsod, R.; Donald, A. M. Rev. Sci. Instrum. 2008, 79, 126107-126101–126107-126103. 43. Rizzieri, R.; Baker, F.; Donald, A. M. Polymer 2003, 44, 5927–5935. 44. Rizzieri, R.; Baker, F.; Donald, A. M. Rev. Sci. Instrum. 2003, 74, 4423–4428. 45. Rizzieri, R.; Mahadevan, L.; Vaziri, A.; Donald, A. M. Langmuir 2006, 22, 3622–3626. 46. Wei, Q. F.; Liu, Y.; Wang, X. Q.; Huang, F. L. Polym. Testing 2007, 26, 2–8. 47. Rawal, A.; Lomov, S.; Verpoest, I. J. Textile Inst. 2008, 99, 235–241. 48. Rinckenbach, S.; Hemmerle, J.; Dieval, F.; et al. J. Biomed. Mater. Res. Part A 2008, 84A, 576–588. 49. Miller, A. F.; Donald, A. M. Biomacromolecules 2003, 4, 510–517. 50. Fechner, P. M.; Wartewig, S.; Kiesow, A.; et al. J. Pharm. Pharmacol. 2005, 57, 689–698. 51. Sgriccia, N.; Hawley, M. C.; Misra, M. Compos. Part A: Appl. Sci. Manuf. 2008, 39, 1632–1637. 52. Jahn, A.; Schroder, M. W.; Futing, M.; et al. Spectrochim. Acta Part A 2002, 58, 2271–2279. 53. Bos, H. L.; Donald, A. M. J. Mater. Sci. 1999, 34, 3029–3034. 54. Turkulin, H.; Holzer, L.; Richter, K.; Sell, J. Wood Fiber Sci. 2005, 37, 552–564. 55. Ma, Q.; Rudolph, V. Drying Technol. 2006, 24, 1397–1403. 56. Eder, M.; Stanzl-Tschegg, S.; Burgert, I. Wood Sci. Technol. 2008, 42, 679–689. 57. Vasic, S.; Stanzi-Tschegg, S. Holzforschung 2007, 61, 367–374. 58. Vasic, S.; Stanzl-Tschegg, S. In Computational Methods and Experiments in Materials Characterisation II; Brebbia, C. A., Mammoli, A. A., Eds.; WIT Press, Southampton UK, 2005; Vol. 51, pp 121–130. 59. Fruhmann, K.; Burgert, I.; Stanzl-Tschegg, S. E. Holzforschung 2003, 57, 326–332. 60. Astley, O. M.; Chanliaud, E.; Donald, A. M.; Gidley, M. J. Int. J. Biol. Macromol. 2001, 29, 193–202. 61. Grande, C. J.; Torres, F. G.; Gomez, C. M.; et al. Mater. Sci. Eng. C. 2009, 29, 1098–1104. 62. James, B. Trends Food Sci. Technol. 2009, 20, 114–124. 63. Uwins, P. J. R.; Murray, M.; Gould, R. J. Microscopy Res. Techn. 1993, 25, 412–418. 64. Thiel, B. L.; Donald, A. M. Ann. Bot. 1998, 82, 727–733. 65. Thiel, B. L.; Donald, A. M. J. Text. Stud. 2000, 31, 437–455. 66. Donald, A.; Baker, F.; Smith, A.; Waldron, K. Ann. Bot. 2003, 92, 73–77. 67. Zheng, T.; Waldron, K. W.; Donald, A. M. Planta 2009, 230, 1105–1113. 68. Wang, S. J.; Yu, J. L.; Jin, F. M.; Yu, J. G. Int. J. Biol. Macromol. 2008, 43, 216–220. 69. Edwards, M. A.; Osborne, B. G.; Henry, R. J. J. Cereal Sci. 2008, 48, 180–192. 70. Blennow, A.; Hansen, M.; Schutz, A.; et al. J. Struct. Biol. 2003, 143, 229–241. 71. Roman-Gutierrez, A. D.; Guilbert, S.; Cuq, B. LWT – Food Sci. Technol. 2002, 35, 730–740. 72. Tang, X.; de Rooij, M.; de Jong, L. Scanning 2007, 29, 197–205. 73. Dang, J. M. C.; Copeland, L. J. Sci. Food Agric. 2004, 84, 707–713. 74. James, B. J.; Smith, B. G. LWT – Food Sci. Technol. 2009, 42, 929–937. 75. Noronha, N.; Duggan, E.; Ziegler, G. R.; et al. Food Res. Int. 2008, 41, 472–479.
Structure Characterization in Real Space | Environmental Scanning Electron Microscopy
545
Biographical Sketch Athene Donald is a professor of Experimental Physics at the University of Cambridge, where she has spent most of her career aside from a 4-year spell at Cornell University in the United States of America. Since her PhD, she has utilized a variety of microscopies in her research, and has been working on ESEM for the past 20 years. She is a soft matter and biological physicist who studies polymers of both synthetic and biological origin, with a particular emphasis on starch and proteins. She has been awarded numerous prizes during her career, including the Faraday and Mott Prizes of the Institute of Physics, the Bakerian Lectureship of the Royal Society, and was the Laureate for Europe in 2009 of the L’Oreal/UNESCO for Women in Science awards. She was elected a fellow of the Royal Society in 1999 and a Dame Commander of the British Empire (DBE) in 2010.
Environmental Scanning Electron Microscopy
AM Donald, University of Cambridge, Cambridge, UK © 2012 Elsevier B.V. All rights reserved.
2.21.1 2.21.2 2.21.3 2.21.4 2.21.4.1 2.21.4.2 2.21.5 2.21.6 References
Introduction The Instrument: A Comparison with Conventional SEM Static Experiments Dynamic Experiments Hydration and Dehydration Mechanical Testing Biopolymers and Biofilms Conclusions
2.21.1 Introduction The technique of environmental scanning electron microscopy (ESEM) has been around for nearly 20 years. However, in many ways it is still in its infancy, and applications to polymer science remain relatively few, despite the advantages it may potentially confer. In part this reflects the fact that for so many purposes conventional scanning electron microscopy (CSEM) looks suf ficient and is familiar, widely available, and easy to use (see Table 1 and Reference 1). In part it would appear to have been due to failing marketing strategies within the companies that manufacture the different varieties of the instrument, initi ally implying the instrument was much simpler to use than many researchers found it, and subsequently failing to appreci ate fully its strengths for hydrated samples such as latex dispersions. However, it also has to be recognized that there are limitations of the instrument, as will be described later in this chapter, and some of these, including the fundamental challenge of beam damage and resolution when performing energy-dispersive X-ray (EDX) analysis, are likely always to limit its utility for certain types of sample and problem. Nevertheless, there are many advantages that can usefully be factored in for polymer scientists trying to study a wide range of different situations, and this chapter will seek to identify the special strengths of the technique.
2.21.2 The Instrument: A Comparison with Conventional SEM There are two key differences with regard to a conventional SEM (CSEM): the chamber around the sample does not need to be under high vacuum and, as a direct consequence of this, insulators do not need to be coated with a conductive coating. These key differences lie at the heart of all the potential advan tages of the instrument. The basic design is essentially the same as a CSEM: there is an electron source, which may (as in CSEM) be a simple tungsten filament, a LaB6 filament, or a field emis sion gun (FEG), each with its own advantages of brightness, durability, and cost. This gun operates under the standard conditions of high vacuum, although exactly how high a vacuum is required depends on the type of gun used. The electron beam travels down the column toward the sample chamber. However, in a CSEM this entire path, with lenses to Polymer Science: A Comprehensive Reference, Volume 2
539 539 541 542 542 543 543 544 544
focus the beam, is under the same high vacuum conditions as the gun, whereas this is not the case in the ESEM. Instead, the column has a series of different pressure zones, as shown in Figure 1, each separated from its neighbor by a pressure limit ing aperture (PLA) across which differential pumping operates. As a result, the pressure increases steadily down the column. This permits the last region, which is the sample chamber itself, to contain a significant pressure of gas, up to around 10–15 torr (∼ 2000 Pa). The upper limit for this pressure is set by a number of considerations including the desired resolution, the type of detector used, and the accelerating voltage. The presence of the gas degrades the image quality less than that which might be expected due to scattering of the incident electron beam by the gas molecules before the electrons ever reach the sample. In practice, as long as the working distance (the distance from the bottom of the last aperture to the sample) is kept small, and the pressure in the chamber is not too large, most electrons are not scattered and those that are, are scattered only a few times; this means beam broadening is not too much of a problem and the shape of the incident beam is still strongly peaked above a rather flat background, as shown in Figure 2. There will be some small signal arising from the further reaches of this flat background, but the majority of the signal will be initiated by the sharp peak at the center. The net effect is to introduce what is essentially a background DC signal on the signal of interest. Quoted instrumental resolution may be < 10 nm, but for polymers this is very unlikely to be realized because of the usual problems of electron beam damage. The type of experiments for which the existence of the broad back ground ‘skirt’ is likely to matter is when X-ray microanalysis is attempted.2 It is the presence of the gas around the sample which makes the key difference with CSEM, and which confers ESEM’s many advantages. Frequently – but not necessarily – this gas is water vapor. If it is, it in principle permits a hydrated sample to be maintained in its hydrated state, and this will be discussed further below. For polymers and other insulators, however, the presence of this ‘imaging gas’ confers a particular advantage because of the way it can reduce the usual problem of charging. In a CSEM, this is overcome by coating the surface of the sample, but the disadvantage of so doing is that fine surface features may be obscured. (Modern low-voltage instruments can to some extent overcome this problem of charging.) In the ESEM, coating is not usually required, although many
doi:10.1016/B978-0-444-53349-4.00044-3
539
540
Table 1
Structure Characterization in Real Space | Environmental Scanning Electron Microscopy
Comparison of conventional and environmental SEM CSEM
ESEM
Gun Sample environment
Tungsten, LaB6, or field emission High vacuum
SE Detector BSE detector X-ray detection Imaging of insulators
Everhart-Thornley Yes Yes Need conductive coating, unless working at very low voltages Only after dehydration
Tungsten, LaB6, or field emission Gas (typically water), pressures up to ∼ 2000 Pa with optional high vacuum Gaseous secondary electron detector Not yet very good at discriminating Yes, but rather poor spatial resolution Yes
Imaging of hydrated samples
Yes
Pressure 10–7 torr
Pressure zone Gun chamber
10–6 torr
Upper column
Anode
10–4 torr
EC2
Column
10–1 torr
EC1
10 torr
Specimen
Gun
Projection apertures Column valve Upper pressure limiting aperture Vacuum manifold
Specimen
Lower pressure limiting aperture and integrated detector
Figure 1 Schematic drawing of the ESEM column, showing the different pressure zones. 1 torr ∼ 133 Pa.
(a)
Primary beam
(b)
Figure 2 In the so-called oligo-scattering regime, electrons in the inci dent probe undergo few, if any, collisions. (a) Schematic of the scattering undergone by the incident beam (heavy arrowed line) in this regime. (b) The resultant probe has a sharp intensity peak superimposed on a low background ‘skirt’.
inexperienced users do not appreciate this fact and may still apply such a coating. But in practice the gas molecules in the sample chamber can be ionized by electrons emitted by the sample (secondary electrons (SEs) predominantly, since their
collision cross section with the gas molecules is higher than the more energetic backscattered electrons (BSEs)). As a conse quence of these ionizing collisions, there is a resultant cascade of electron–gas molecule ionizations which is shown schematically in Figure 3. Each ionizing collision produces a positively charged ion together with an additional so-called daughter electron, and this leads to amplification of the electrons traveling toward the detector. The magnitude of this amplification can be very significant.3 However, in the context of charging, what matters is the generation of a substantial number of positive gaseous ions. These tend to drift toward the negatively charged sample – negatively charged because of the implantation of incident electrons – and as they accumulate at the surface, they tend to compensate for the negative charge buildup, and so reduce the effect of charging. The processes involved are complex, and the interested reader should look at the review4 for more details. In general, however, the effect means that insulators do not need a conductive coating. Furthermore, if a new surface is opened up, for instance during a dynamic experiment to study fracture, charging is still not an issue; examples of this will be given below. In general, this means that ESEM is particularly powerful for dynamic
Structure Characterization in Real Space | Environmental Scanning Electron Microscopy
541
Positively charged detector Electron beam Cascade electrons
Vo
A
Positivion
+ Gaseous atom V
Earthed sample Figure 3 Schematic representation of the cascade amplification process that operates when a gas is present in the sample chamber. Electrons emitted from the sample collide with gaseous atoms and can ionize them, producing additional daughter electrons that contribute to the signal detected at the positively charged detector.
experiments of insulators. In addition, the absence of a coating means that the intrinsic contrast from the sample is not obscured as the electrons emerge from the sample surface: for polymers, with their typical low atomic numbers and small differences between chemically distinct species, this can confer a further advantage that has been little exploited in the literature. So there are two fundamental advantages of the ESEM over CSEM: the sample can be imaged in a hydrated state, and insulators do not need to be coated. Let us now look at the signal that is detected. In CSEM, the detector works by having a positive bias to which the electrons are attracted. Conventionally, there are two detectors for detecting the low-energy SEs, usually defined as those with energies < 50 eV, and the high-energy BSEs. The latter are electrons deflected through close to 180° on hitting the sample, whereas the former are electrons emitted from the sample due to inelas tic processes (ionization, etc.). Because the SEs possess low energy, they can only escape from close to the sample surface, and, therefore, are particularly useful at providing topographic information. In contrast, BSEs can come from much deeper within the sample, and the number generated depends on atomic number. Thus BSEs typically can provide information on chemical composition. In CSEM, the two detectors are set up to distinguish the different signals, and, therefore, topo graphic and chemical information can be obtained separately. However, these standard detectors do not work in the low vacuum conditions of an ESEM, and existing detectors for the ESEM are much less successful at distinguishing the different signals. An additional problem arises because of the nature of
the cascade amplification itself. This process amplifies all signals, thus amplifying noise such as that generated by BSEs hitting other parts of the chamber and causing additional electrons to be generated which contain no useful information. These electrons are known as SE2s, and tend simply to degrade image quality. A discussion of signal composition can be found in Reference 5. In practice, good BSE detectors are still not available for the ESEM, and the SE detector will not completely discriminate between topographic and compositional contrast. Finally, the ever-present problem of electron beam damage should be discussed. The origins of beam damage for polymers in conventional microscopy have been amply discussed in previous articles.6,7 For ESEM, there are some additional issues that need to be borne in mind, notably that there can be additional damage mechanisms, particularly in the presence of water. Relatively little work has been done specifically study ing these effects,8 but the work that has been done has demonstrated that beam damage is worse because of the free radicals generated by collisions between the water molecules and the electrons,9,10 and these free radicals can diffuse across the sample surface causing additional damage to that directly caused by the incident electrons.9,10
2.21.3 Static Experiments As indicated above, the presence of the gas in the chamber means that typically charging is no longer an issue for insula tors, as in CSEM, so that these samples can be studied without a conducting coating obscuring surface detail. Nevertheless, there
542
Structure Characterization in Real Space | Environmental Scanning Electron Microscopy
is no doubt that the presence of a thin gold layer may ‘sharpen’ the image, and it is presumably for this reason that some papers still adopt this approach.11,12 However, for other applications, the absence of a coating may reveal contrast otherwise obscured by it. This was initially observed for a nonpolymeric sample of water–oil emulsions.13 The origin of the contrast lies in the differences in electronic properties between the two components, and the consequent mechanisms by which electrons traveling to the surface can lose energy. Similar con trast effects have also been observed in ceramics such as gibbsite,14 although with a subtly different interpretation. For polymers, this has particular advantages because differences in atomic number are usually tiny, although some attempts have been made to explore chemical differences with BSEs.15 For instance, in block copolymers of polystyrene-polyisoprene, only carbon and hydrogen are pre sent just arranged in different ways and with slightly different densities. Nevertheless, clear contrast can be seen between the phases (Figure 4) without the customary use of staining to reveal the microstructure, for instance via the use of osmium tetroxide. Indeed, recent work is indicating that the use of such stains may actually preferentially swell one of the phases, meaning that measurement of lamellae spacings is inaccu rate,16 thus highlighting the particular value the minimal sample preparation required for imaging in the ESEM can bring. However, since the source of the contrast is related to the energy loss processes that occur as the electrons travel from their site of generation to the sample surface, the thickness of the material (for a thin film) through which the electrons must
2 µm
Figure 4 Contrast is visible without staining in a film of polystyrene-polyisoprene block copolymer with 50% of each component present. The polystyrene lamellae appear darker than the polyisoprene due to density differences. Micrograph courtesy of AJ Ryan and C Salou (University of Sheffield).
travel will play a part. So where there is significant topography present, care must be taken in order to interpret the images correctly. This has been demonstrated for the case of semicon ducting polymer blends spin cast from solution, in which significant topography is introduced during the drying process. Strong contrast is seen, but thought must be given as to which phase is identified with which polymer.17
2.21.4 Dynamic Experiments 2.21.4.1
Hydration and Dehydration
The ability to maintain a sample in a hydrated state – or indeed saturated with some fluid other than water – has value for many polymer applications, including those relevant to filtra tion and textiles. These appear to be the fields – other than biopolymer applications discussed further below – which have received most attention, but limited work has been carried out where a conventional polymer is explored in a hydrated state.18 The ability to change the level of hydration – either by deposit ing water on a sample or by dehydrating the sample and following the consequent morphological changes – opens up even more avenues. This can be done by changing the water vapor pressure in the chamber. Figure 5 shows the variation of the saturated vapor pressure (SVP) for water as a function of temperature, and indicates how small variations in temperature can shift the sample from hydrating to dehydrating conditions. Typically, the work needs to be done below room temperature, otherwise the requisite pressures are too high for useful ima ging, as can be inferred from Figure 5, but this does not alter the basic utility of this approach. For textiles, this has led to studies of swelling19–22 and analysis of the shape of water droplets21 and contact angle measurement of the droplets on the fibers.23,24 Although it might be thought that optical ima ging provided quite enough resolution to enable the contact angle to be measured accurately on a textile fiber, in practice Fresnel fringes often obscure the actual water–fiber interface, rendering precise measurement difficult, plus in principle local heterogeneities – leading to local variations in contact angle – can be explored, although this route does not seem to have been exploited. For membranes, some work has fully exploited the capabil ities of the ESEM to carry out dynamic experiments. For instance, as with the fiber work described above, Yu et al.25 have used the ability both to condense water droplets onto membranes to characterize their hydrophilicity/hydrophobi city and to explore the resulting contact angles. Wang et al.26 controlled the chamber relative humidity to permit hydration/ dehydration experiments to be carried out while observing the porosity of the membranes. Le-Clech et al.27 compared differ ent microscopic approaches for the study of biofouling of membranes, a very important problem in many applications, and noted that, although ESEM provided a major advance in being able to image the samples in appropriate wet mode, it was hard to focus on the deposited (alginate) films to observe fine structure. This will always be a challenge for ESEM, so it is a technique that most usefully works in tandem with other approaches in a complementary way. Cryogel structure during drying was also studied for polyacrylamide gels with different degrees of cross-linking so that the pore structure could be clearly visualized.28
Structure Characterization in Real Space | Environmental Scanning Electron Microscopy
543
25
Pressure (Torr)
20
Liquid
15 Condense
Evaporate
10 5 0
Gas
0
5
10
15
20
25
Temperature (°C) Figure 5 The SVP curve for water vapor, indicating the regimes in which evaporation or condensation will occur.
Additionally, solvents other than pure water can be used. This was done by Zhong et al. looking at aggregation of a terpolymer based on acrylamide in solutions of pure water and with salt present29 and with colloidal PMMA,30 in which the effect of the modification of the electrostatic screening due to the presence of salt was studied. This latter study highlights one particular class of polymeric samples, that of latices, that have been extensively studied during dehydration in the ESEM to reveal the stages in morphological development including the kind that make up paint formulations. The first researcher to carry out such experiments was Eckersley,31 and since then a range of different formulations have been explored, and novel insight obtained on the film drying process.32–37 In particular, a combination of ESEM with ellipsometry allowed the identifica tion of an intermediate stage in the standard stages of the drying process, in which most of the interstitial water had been lost, but complete densification was not achieved.38 Another variant of ESEM has also been used to study latex formulations by Bogner’s group. In this case, a wet-STEM (scan ning transmission electron microscopy) detector was used, so that the image was formed below the sample and the aggrega tion was observed.39
2.21.4.2
Mechanical Testing
As mentioned above, the relaxation of the condition for coating a sample surface means that ESEM offers particular attractions for performing mechanical tests in situ. Despite this, many of the papers that utilize ESEM in the context of mechanical testing only use the technique to study postmortem fracture surfaces. Rather few groups working on polymers have estab lished the requisite testing rigs. In general, these will need a cooling module as well, for the reasons given above about the useful SVP curves for water, but also because for polymers with glass transition temperatures close to room temperature, accu rate temperature control is essential if meaningful results are to be obtained.40 Different testing geometries can also be con structed: for instance Nase et al. have demonstrated the ability to carry out in situ peel tests on polymer bilayers,41 and Dragnevski et al. have produced a rig that allows for large extensions of up to more than 100% to be achieved.42
But the bulk of the work in the literature has used simple tensile/compression tests, allowing correlation of the deforma tion with the load-extension curves.43–48 In this way, it is possible to identify the nature of the failure route, or the loca tion of first yield in a heterogeneous sample. It would seem there is ample scope to extend the fairly limited studies that have been carried out to date.
2.21.5 Biopolymers and Biofilms The examples chosen so far have been focused on synthetic polymers of different types. In fact, ESEM has been particularly widely applied to biopolymers and biologically derived sam ples including biofilms, tissues, and bacteria, all of which contain substantial polymeric components. Cellulose has been explored in various states, both native fibers and wood and also dispersions of cellulose microfibrils. For all these types, the ability ‘not’ to dehydrate the sample has proved beneficial. In the case of microfibrillar assembly, how ever, it has also been the case that studying the arrangements of the microfibrils in dispersion49 and the way they aggregate has been possible at quite high resolution even in water. Many natural fibers have been studied ranging from hemp,50,51 through flax,52 including in situ mechanical testing,53 to wood. For the latter, there have been extensive studies on optimization of imaging54 and in situ swelling,55 and most notably, micromechanical studies.56–59 That the wood does not need to be dried or coated has clearly been identified as a key advantage for these studies. Composites containing cellu lose fibers have also been studied via ESEM, notably involving bacterial (rather than plant) produced cellulose.60,61 Food – broadly defined – is another area in which the ability to study wet systems has proved very useful. A recent review by James62 covers work in the field. Food here covers both vegetables (which could also be included in the cellulose section above) and starch-based materials, but also systems containing proteins and fats, although the last two categories have been much less exten sively studied than the former. The study of vegetables in the ESEM goes back to Uwins in 1993,63 who compared different sample preparation routes for potatoes, including the minimal preparation required for ESEM. In situ deformation of carrots
544
Structure Characterization in Real Space | Environmental Scanning Electron Microscopy
under different geometries64,65 and, more recently, onions66,67 has been used to look at failure modes correlated with load-extension data. Starch-based and starch-containing foods have been studied with various different motivations ranging from simply cataloging samples that had not been extensively treated prior to observation,68–70 to following hydration in situ by changing the sample chamber pressure,71,72 to carrying out in situ fracture tests.73 Rather less work has been done on other types of systems, although both fat-containing74,75 and fat-replacement systems43 have been looked at.
2.21.6 Conclusions ESEM offers many advantages for the polymer scientists, most notably the ability to image samples without applying a con ductive coating and the possibility of carrying out dynamic experiments – in situ hydration, dehydration, or mechanical tests. However, only certain parts of the community appear to have appreciated the potential of the technique fully, and there are many other types of sample upon which few ESEM experiments have been carried out. There are undoubtedly limitations, including beam damage (particularly in the pre sence of water vapor and the consequent generation of damaging free radicals) and, by not applying a coating, images may frequently look less sharp as if the resolution is worse than it truly is. Nevertheless, there are many advantages which the technique can confer, and it is to be hoped it will find an increasingly important place in the armory of characterization techniques used by the polymer science community.
References 1. Goldstein, J. I.; Newbury, D. E.; Joy, D. C.; et al. Scanning Electron Microscopy and X-ray Microanalysis, 3rd ed.; Springer: New York, 2002. 2. Mansfield, J. F. Mikrochim. Acta 2000, 132, 137–143. 3. Thiel, B.; Bache, I. C.; Fletcher, A. L.; et al. J. Micros. 1997, 187, 143–157. 4. Thiel, B. L.; Toth, M. J. Appl. Phys. 2005, 97, 051101. 5. Fletcher, A. L.; Thiel, B. L.; Donald, A. M. J. Micros. 1999, 196, 26–34. 6. Tsuji, M. In Comprehensive Polymer Science; Allen, G., Bevington, J. C., Eds.; Pergamon: Oxford, 1989; Vol. 1, p 803. 7. Sawyer, L. C.; Grubb, D. T.; Meyers, G. F. Polymer Microscopy, 3rd ed.; Springer: New York, 2008. 8. Zankel, A.; Chernev, B.; Brandl, C.; et al. Macromol. Symp. 2008, 265, 156–165. 9. Kitching, S.; Donald, A. M. J. Micros. 1998, 190, 357–365. 10. Royall, C. P.; Thiel, B. L.; Donald, A. M. J. Micros. 2001, 204, 185–195. 11. Sun, J.; Chen, X. S.; Lu, T. C.; et al. Langmuir 2008, 24, 10099–10106. 12. Guo, Z. P.; Li, Y. H.; Tian, H. Y.; et al. Langmuir 2009, 25, 9690–9696. 13. Stokes, D. J.; Thiel, B. L.; Donald, A. M. Langmuir 1998, 14, 4402–4408. 14. Watt, G. R.; Griffin, B. J.; Kinny, P. D. Am. Mineral. 2000, 85, 1784–1794. 15. Franz, N.; Ahlers, M. O.; Abdullah, A.; Hohenberg, H. J. Mater. Sci. 2006, 41, 4561–4567. 16. EMAG. J. Phys. Conf. Series. 2009, 241, (012077). 17. Williams, S. J.; Donald, A. M.; Thiel, B. L.; Morrison, D. E. Scanning 2005, 27, 190–198. 18. Miller, A. F.; Cooper, S. J. Langmuir 2002, 18, 1310–1317. 19. Jenkins, L. M.; Donald, A. M. Scanning 1997, 19, 92–97. 20. Jenkins, L. M.; Donald, A. M. Textile Res. J. 2000, 70, 269–276. 21. He, C. C.; Gu, Z. Y.; Tian, J. Y. J. Appl. Polym. Sci. 2003, 89, 3939–3943. 22. Wei, Q. F.; Wang, X. Q.; Mather, R. R.; Fotheringham, A. F. Fibres Text. East. Eur. 2004, 12, 79–83. 23. Jenkins, L. M.; Donald, A. M. Langmuir 1999, 15, 7829–7835. 24. Wei, Q. F.; Mather, R. R.; Fotheringham, A. F.; Yang, R. D. Textile Res. J. 2003, 73, 557–561.
25. Yu, H. M.; Schumacher, J. O.; Zobel, M.; Hebling, C. J. Power Sources 2005, 145, 216–222. 26. Wang, J.; Dismer, F.; Hubbuch, J.; Ulbricht, M. J. Membr. Sci. 2008, 320, 456–467. 27. Le-Clech, P.; Marselina, Y.; Ye, Y.; et al. J. Membr. Sci. 2007, 290, 36–45. 28. Plieva, F. M.; Karlsson, M.; Aguilar, M. R.; et al. Soft Matter 2005, 1, 303–309. 29. Zhong, C. R.; Ye, L.; Dai, H.; Huang, R. H. J. Appl. Polym. Sci. 2007, 103, 277–286. 30. He, C.; Donald, A. M. Langmuir 1996, 12, 6250–6256. 31. Eckersley, S. T.; Rudin, A. Prog. Org. Coat. 1994, 23, 387–402. 32. Keddie, J.; Meredith, P.; Jones, R.; Donald, A. In ACS Symposium Series; Provder, T., Winnik, M., Urban, M., Eds.; ACS: Washington, DC, 1996; Vol. 648, pp 332–348. 33. Meredith, P.; Donald, A. M. J. Micros. 1996, 181, 23–35. 34. Donald, A. M.; He, C.; Royall, C. P.; et al. Colloids Surf., A 2000, 174, 37–53. 35. Royall, C. P.; Donald, A. M. Scanning 2002, 24, 305–313. 36. Kugge, C.; Craig, V. S. J.; Daicic, J. Colloids Surf., A: Physicochem. Eng. Aspects 2004, 238, 1–11. 37. Dragnevski, K. I.; Donald, A. M. Colloids Surf., A: Physicochem. Eng. Aspects 2008, 317, 551–556. 38. Keddie, J. L.; Meredith, P.; Jones, R. A. L.; Donald, A. M. Macromolecules 1995, 28, 2673–2682. 39. Bogner, A.; Guimaraes, A.; Guimaraes, R. C. O.; et al. Colloid. Polym. Sci. 2008, 286, 1049–1059. 40. Zankel, A.; Poelt, P.; Gahleitner, M.; et al. Scanning 2007, 29, 261–269. 41. Nase, M.; Zankel, A.; Langer, B.; et al. Polymer 2008, 49, 5458–5466. 42. Dragnevski, K. I.; Fairhead, T. W.; Balsod, R.; Donald, A. M. Rev. Sci. Instrum. 2008, 79, 126107-126101–126107-126103. 43. Rizzieri, R.; Baker, F.; Donald, A. M. Polymer 2003, 44, 5927–5935. 44. Rizzieri, R.; Baker, F.; Donald, A. M. Rev. Sci. Instrum. 2003, 74, 4423–4428. 45. Rizzieri, R.; Mahadevan, L.; Vaziri, A.; Donald, A. M. Langmuir 2006, 22, 3622–3626. 46. Wei, Q. F.; Liu, Y.; Wang, X. Q.; Huang, F. L. Polym. Testing 2007, 26, 2–8. 47. Rawal, A.; Lomov, S.; Verpoest, I. J. Textile Inst. 2008, 99, 235–241. 48. Rinckenbach, S.; Hemmerle, J.; Dieval, F.; et al. J. Biomed. Mater. Res. Part A 2008, 84A, 576–588. 49. Miller, A. F.; Donald, A. M. Biomacromolecules 2003, 4, 510–517. 50. Fechner, P. M.; Wartewig, S.; Kiesow, A.; et al. J. Pharm. Pharmacol. 2005, 57, 689–698. 51. Sgriccia, N.; Hawley, M. C.; Misra, M. Compos. Part A: Appl. Sci. Manuf. 2008, 39, 1632–1637. 52. Jahn, A.; Schroder, M. W.; Futing, M.; et al. Spectrochim. Acta Part A 2002, 58, 2271–2279. 53. Bos, H. L.; Donald, A. M. J. Mater. Sci. 1999, 34, 3029–3034. 54. Turkulin, H.; Holzer, L.; Richter, K.; Sell, J. Wood Fiber Sci. 2005, 37, 552–564. 55. Ma, Q.; Rudolph, V. Drying Technol. 2006, 24, 1397–1403. 56. Eder, M.; Stanzl-Tschegg, S.; Burgert, I. Wood Sci. Technol. 2008, 42, 679–689. 57. Vasic, S.; Stanzi-Tschegg, S. Holzforschung 2007, 61, 367–374. 58. Vasic, S.; Stanzl-Tschegg, S. In Computational Methods and Experiments in Materials Characterisation II; Brebbia, C. A., Mammoli, A. A., Eds.; WIT Press, Southampton UK, 2005; Vol. 51, pp 121–130. 59. Fruhmann, K.; Burgert, I.; Stanzl-Tschegg, S. E. Holzforschung 2003, 57, 326–332. 60. Astley, O. M.; Chanliaud, E.; Donald, A. M.; Gidley, M. J. Int. J. Biol. Macromol. 2001, 29, 193–202. 61. Grande, C. J.; Torres, F. G.; Gomez, C. M.; et al. Mater. Sci. Eng. C. 2009, 29, 1098–1104. 62. James, B. Trends Food Sci. Technol. 2009, 20, 114–124. 63. Uwins, P. J. R.; Murray, M.; Gould, R. J. Microscopy Res. Techn. 1993, 25, 412–418. 64. Thiel, B. L.; Donald, A. M. Ann. Bot. 1998, 82, 727–733. 65. Thiel, B. L.; Donald, A. M. J. Text. Stud. 2000, 31, 437–455. 66. Donald, A.; Baker, F.; Smith, A.; Waldron, K. Ann. Bot. 2003, 92, 73–77. 67. Zheng, T.; Waldron, K. W.; Donald, A. M. Planta 2009, 230, 1105–1113. 68. Wang, S. J.; Yu, J. L.; Jin, F. M.; Yu, J. G. Int. J. Biol. Macromol. 2008, 43, 216–220. 69. Edwards, M. A.; Osborne, B. G.; Henry, R. J. J. Cereal Sci. 2008, 48, 180–192. 70. Blennow, A.; Hansen, M.; Schutz, A.; et al. J. Struct. Biol. 2003, 143, 229–241. 71. Roman-Gutierrez, A. D.; Guilbert, S.; Cuq, B. LWT – Food Sci. Technol. 2002, 35, 730–740. 72. Tang, X.; de Rooij, M.; de Jong, L. Scanning 2007, 29, 197–205. 73. Dang, J. M. C.; Copeland, L. J. Sci. Food Agric. 2004, 84, 707–713. 74. James, B. J.; Smith, B. G. LWT – Food Sci. Technol. 2009, 42, 929–937. 75. Noronha, N.; Duggan, E.; Ziegler, G. R.; et al. Food Res. Int. 2008, 41, 472–479.
Structure Characterization in Real Space | Environmental Scanning Electron Microscopy
545
Biographical Sketch Athene Donald is a professor of Experimental Physics at the University of Cambridge, where she has spent most of her career aside from a 4-year spell at Cornell University in the United States of America. Since her PhD, she has utilized a variety of microscopies in her research, and has been working on ESEM for the past 20 years. She is a soft matter and biological physicist who studies polymers of both synthetic and biological origin, with a particular emphasis on starch and proteins. She has been awarded numerous prizes during her career, including the Faraday and Mott Prizes of the Institute of Physics, the Bakerian Lectureship of the Royal Society, and was the Laureate for Europe in 2009 of the L’Oreal/UNESCO for Women in Science awards. She was elected a fellow of the Royal Society in 1999 and a Dame Commander of the British Empire (DBE) in 2010.