The electron source is the most important component of the Scanning Electron Microscope (SEM) since it is this which will determine the overall performance of the machine. The gun performance can be described in terms of quantities such as its brightness, its source size, its energy spread, and its stability and, depending on the chosen application, any of these factors may be the most significant one. The task of the electron gun in a SEM is, in fact, particularly difficult because of the very wide range of operational parameters that may be required, e.g. a variation in probe size of from a few angstroms to a few microns, and a probe current which may go from less than a pico-amp. to more than a microamp. This wide range of operating parameters makes the choice of the optimum source for scanning microscopy a difficult decision.

The study of plant morphology and plant cells in the scanning electron microscope is often compromised by the limitations of specimen preparation techniques. Air drying usually results in unacceptable shrinkage and distortion of the normal surface morphology of plant cells. Chemical fixation followed by critical point drying or chemical drying using fluorocarbon compounds improves morphological results but still imparts artifacts, adds chemical constituents to the specimen, and requires the use of toxic chemicals, a hood, and much time.

One technique that eliminates many of these disadvantages and is even suitable for specimen preparation in the field is tissue printing. For easy, quick recording of stem anatomy and collection of cell exudates for subsequent analysis, its “elegant simlicity” is compelling. Historically, the application of tissue printing has been in connection with optical microscopy. However, this technique works very well for SEM and associated elemental characterization of residues by x-ray microanalysis. The tissue print technique applied to SEM is much the same as for optical microscopy.

In today's FTIR market, many instrument manufacturers provide software to perform searches of computer library spectra. With the increased speed of computers running FTIR instruments, you can have the result of a computer library search in literally seconds. However, as we rely more and more on computers in the laboratory we need to keep in mind the limitations and possible areas for error in these applications.

Computer library search programs calculate the difference between the sample spectrum and reference spectrum typically based on peak position and intensity. Different searching algorithms can stress one or the other. The result of a computer search is a Hit List with each hit having a hit quotient index (HQI) to indicate how close that particular spectrum matches the sample spectrum. In some programs the quotient which indicates an exact match would be 1 or 100, in others it is 0.

A combination of electron microscope and x-ray spectrometer is a very powerful tool. Not only can one see a sample in great detail, but one can determine, and even map, the chemical elements. In Part 1, I discussed some of the basics of energy dispersive x-ray spectroscopy (EDS or EDX). The heart of the spectrometer is a small piece of single crystal silicon about the size and shape of a shirt button, and about twice as thick. It has been selected for high purity, and then lithium drifted to compensate the remaining impurities.

The lithium is carefully drifted into the crystal button in order to exactly compensate the impurities in the crystal that would create leakage current. This is done at about 60° C under an electric field. It is then evaluated and re-drifted for a final clean up of any uncompensated impurity atoms that remain.

At present most medical microprobe analysis is conducted on insoluble particulates such as asbestos fibers in lung tissue. Cryotechniques are not necessary for this type of specimen. Insoluble particulates can be processed conventionally Nevertheless, it is important to emphasize that conventional processing is unacceptable for specimens in which electrolyte distributions in tissues are sought. It is necessary to flash-freeze in order to preserve the integrity of electrolyte distributions at the subcellular and cellular level. Ideally, biopsies should be flash-frozen in the operating room rather than being frozen several minutes later in a histology laboratory. Electrolytes will move during such a long delay. While flammable cryogens such as propane obviously cannot be used in an operating room, liquid nitrogen-cooled slam-freezing devices or guns may be permitted, and are the best way to achieve an artifact-free, accurate tissue sample which truly reflects the in vivo state. Unfortunately, the importance of cryofixation is often not understood.

No, Virginia, there is no objective lens in the Scanning Electron Microscope.

There is much confusion about the lenses in a scanning electron microscope (SEM). All of the lenses in a SEM are condenser lenses. The fact that some scanning electron microscope manufacturers have labelled the controls for the final condenser “objective lens” is misleading and this may have contributed to the confusion.

To get some perspective on this matter, observe your compound light microscope. Starting with the light source, the first lens encountered is the condenser lens (the lens that handles the light before it interacts with the specimen). Next the light interacts with the specimen on a microscope slide. Then the light enters the objective lens (the first lens the light enters after interacting with the specimen) and finally the light travels through the ocular lens and then it enters your eye.

Recent advances in digital image processing hardware providing very high pixel resolutions can be exploited to solve problems in correlating high magnification feature analysis with specimen macrostructure. In neuroscience, for instance, analyses such as neuron counting which are performed at high magnification often need to be correlated with the overall structure of the specimen. Typically these procedures involve tediously moving between very low and very high power objectives. Once at high power, the positional knowledge of the objects of interest can be difficult to maintain.

By integrating high resolution image capture with precise microscope automation, a digital composite image of the specimen can be created. Such composite images show the researcher a high resolution “map” of the specimen, facilitating the analysis of both macro- and microstructure characteristics from a single view.

LaB6 (and CeB6) cathodes are widely used as high-brightness cathodes in a variety of electron beam applications. These cathodes provide about 10 times the brightness and about 100 times the service life of a tungsten filament. However, to fully realize the improved brightness and lifetime of the LaB6 (CeB6) cathode, it must be designed and manufactured specifically for the application.

This article covers tip shape and crystal orientation. Future articles will deal with materials selection, preparation, purity, composition, and cathode operation.

The design of the LaB6 cathode tip is critical for maximum lifetime and optimum performance. The cathode must also provide the appropriate source size, beam current, and beam brightness for the application.

The lifetime of the LaB6 cathode is limited by three basic factors: cathode temperature, vacuum pressure, and tip shape. The first two factors are critical and depend on proper user operation. Poor vacuum and excessive temperatures accelerate LaB6 material loss, which decreases the cathode lifetime. The third factor, tip shape, is also critical, and careful production control ensures that the tip shape is optimized.

Microscopes are known for demonstrating structure, but there have been many examples of how this is changing. A novel use of the atomic force microscope (AFM) by Manfred Radmacher, Monika Fritz, Helen Hansma, and Paul Hansma at the University of California, Santa Barbara, has extended the versatility of this instrument even further. They demonstrated a conformational change in a molecule that happened in a fraction of a second.

Radmacher et al. pointed out that enzymes perform their biologic function by undergoing a conformational change and that this change is within the spatial and temporal resolving capabilities of the AFM. They set out to prove this by lowering the tip of the AFM onto lysozyme, a molecule that is roughly egg-shaped, measuring about 4.5 nm by 3 nm by 3 nm. Using the tapping mode in liquid, they found that single molecules exhibited a spike-like height fluctation when in the presence of a suitable substrate.

X-ray microanalysis (EDS) on a SEM or TEM is a valuable tool for research and industry. No other technique can provide so much information so easily about the chemistry of micro-volumes of a material. Many samples require nothing more than positioning of the electron beam onto the area of interest, collecting a spectra, and 30 seconds later you have your answer. Or do you?

Any technique as widespread and straightforward as energy dispersive x-ray spectroscopy is bound to be misused on occasion. It has been my experience that many users either overestimate the capabilities of EDS to answer difficult materials questions or do not fully appreciate the limitations of the technique and how to work around them. These problems tend to show up as the need to analyze increasingly complex samples increases. The following brief description of the physics of electron beam/specimen interaction may be useful in explaining why problems can exist.

Microscopists often recrystallize or precipitate compounds for purposes of identification. Almost as often, a few crystals form that are irresistibly beautiful and demand isolating for single-crystal x-ray diffraction, spindle stage, polarized IR absorption, or remounting in a crystal-rolling medium like Aroclor® 1260.

The isolation of one such crystal among hundreds from the center of a drop is not as difficult as it sounds. It is done at 50-100X under a stereomicroscope using a fine, usually a tungsten, needle. The needle may (rarely) need to be surface-treated by rubbing with a water-insoluble wax (then tissue-wiped “clean”) to prevent creeping of the solution up the needle and causing solution (and crystal) movement.

The needle is used to clear a path for the desired crystal by pushing the edge crust and other intervening crystals to one side. Complete clearance is not necessary. The desired crystal is then pushed with the needle along the slide to the edge and then well outside of the drop. A few smaller crystals and much solution may accompany it.

In the early days of biological electron microscopy (the 1950s and 1960s), projection slides of electron micrographs for talks or teaching were generally prepared as 3-1/2″ X 4″ lantern slides, which were shown using large lantern-slide projectors. It was felt by professional electron microscopists that the detail, tones, and general image quality of electron micrographs could be adequately portrayed only in this larger format.

However the large lantern slides were very cumbersome, and most professionals began switching to 2″ X 2″ slides in the early 1970s. Some of us during that period put in a great deal of time trying to work out a procedure for printing EM negatives directly on 2″ X 2″ glass Kodak Projector Slide Plates, following the general approach by which we had previously made the 3-1/4″ X 4″ lantern slides.

Having been brought up on monocular microscopes I find the omnipresent binocular systems a luxury. To support this viewpoint I'd like to suggest some benefits you may not have considered.

Because I'm used to monocular viewing I sometimes use two different oculars, say 10X and 25X, in order to scan quickly to find an area of interest and then to examine the detail with higher magnification. Occasionally I use both oculars simultaneously and “concentrate” on either image to the exclusion of the other. A better way is to set the interocular distance at the extreme setting most different from your own interocular distance. By moving your head about a centimeter either way you can use either ocular.

A variation of this theme is very helpful when you are trying to find good crystals showing useful interference figures.

The Confocal microscopy listserver discussion group was put on the air over 5 years ago by Don Parsons (Albany), Steve Paddock (Wisconsin), P.C. Cheng (Buffalo) and me at the suggestion of Parsons that there should be an open discussion forum for this rapidly developing technology. Parsons is an expert in infomatics (among other things) and appreciated the potential of the “mail reflector” concept at an early stage in its development.

Listserver groups were pretty uncommon then but we figured that since we were dealing with digital microscopists, most of those interested would have computers and access to Bitnet or Internet. For the first month or so Parsons, Paddock, Cheng and I corresponded with one another to work wrinkles out of the system and we then began to publicize the existence of the group at meetings, symposia and through the commercial manufacturers of confocal microscopes and associated hardware.

The Shared Research Equipment (SHaRE) Program at Oak Ridge National Laboratory (ORNL) provides microanalytical facilities to researchers from U.S. universities, industries, and government laboratories for studies within the materials sciences. SHaRE is one of four microscopy and microanalysis research user facilities supported by the U.S. Department of Energy's Office of Basic Energy Sciences (BES). All BES user facilities, including SHaRE, are available to in order to foster collaborative research, scientific leadership, international competitiveness, and the education of graduate students being prepared for careers in materials science. Based on the scientific excellence and relevance of the proposed experiments, free operating time is available to researchers who then publish their results in open literature. Proprietary research can be accomplished on a full cost-recovery basis or by Cooperative Research and Development Agreements (CRADAs). However, services that can be purchased from commercial laboratories are not appropriate for the SHaRE Program.

There is a gap between biochemical findings and ultrastructural interpretation of “glycogen granules”. Biochemists have recognized that glycogen contains covalently bound proteins. These include enzymes involved in giycogen metabolism: glycogenin (protein primer responsible for initiation of glycogen synthesis), glycogen synthase and phosphorylase, and presumably other regulatory enzymes. The structures formed by the association of glycogen and protein have been called protein-glycogen complexes, considered as proteoglycans, or as dynamic cellular organelles, glycosomes.

The question arises as to why the biochemical recognition of a protein component in glycosomes has not been acknowledged in electron microscopy (EM)? This protein is visible in every section stained by uranium (U) and lead (Pb) salts where it appears as 20-30 nm granules (Fig. 1). However, these granules are commonly interpreted as glycogen, despite the fact that glycogen does not react with ionic compounds and therefore cannot be stained by U-Pb.

EDS analysis of samples in a SEM or TEM are usually portrayed as a very simple process; just image your specimen, turn on the x-ray analyzer and all will be revealed about the chemistry of your sample. Unfortunately, that approach can be very misleading at best, and disastrous at worst. The analyst should get used to preparing himself for EDS x-ray analysis just like he or she does for any other process. A set of preparation steps should be followed until they become an unconscious method to be used before accepting the information coming from the EOS x-ray spectrum. After proper basic preparation, interpretation of the results and the selection of operating parameters is still very important due to the detailed nature of the specimen.

Preparation: The most basic need is for the EDS system to be calibrated. This can be a difficult or simple task depending on the age of the system.