There are many transmission electron microscopes (TEMs) that have provided years of reliable service and good images, but they lack the positioning systems that are standard on the newest microscopes. A precise digital positioning capability is essential for many quantitative stereological studies and relocating specific features on a grid. For many of the older TEMs, retrofitting them with a digital positioning system is not an option, or at least a very expensive upgrade. Now Martin Hohmann-Marriott, William Sharp, Robert Roberson, and Robert Blankenship have described the hardware and software that offer a second life to these older microscopes.

Over the past few years there has been a dramatic improvement in microscopy acquisition techniques, in effective imaging modalities as well as raw hardware performance. As the microscopist's available tools become more sophisticated and diverse—e.g., time-lapse, Z sectioning, multispectra, lifetime, nth harmonic, polarization, and many combinations thereof—we face a corresponding increase in complexity in the software for understanding and interpreting the resultant data. With lifetime imaging, for example, it is overwhelming to study the raw numbers; instead, an exponential curve-fitting algorithm must be applied to extract meaningful lifetime values from the mass of photon counts recorded by the instrument.

With the recent success of the human genome project, researchers must now discover biological functions of the tens of thousands of proteins encoded in the genome. Molecular cell biology and structural biology methods are required for this work.

Modern multidimensional bioimaging methods, such as laser scanning confocal microscopy and other light microscopic techniques, have emerged as a major research methodology in this new era of informational biology. These methods all produce vast amounts of digital information and require specialized computer software to process and analyze it. Several programs intended for this purpose have become available in recent years, but microscopists around the world frequently discover them unsuitable for many of their required tasks. The reasons for this are varied, and include: extensive bugs that take too long to fix, prohibitively high prices (also for updates and bug fixes), algorithms that are undisclosed and as such unacceptable for accurate scientific work, limited sets of features dispersed into separate “modules”, complex and difficult user interfaces, and lack of flexibility in adapting to specific research situations.

With the mapping of the mouse genome, the growing sophistication in transgenic sciences creating complex mouse models of disease, and the demand to study disease in vivo, there has been a corresponding increase in the demand for and development of preclinical imaging modalities. Clinical ultrasound operating in the 2-12 MHz range is a well established clinical imaging modality, accounting for more than one-third of all imaging procedures performed in North America. The simplicity, ease of use, speed, and safety of ultrasound have led to its significant role in diagnosis, treatment assessment, follow-up, and guidance of therapy in clinical applications. Ultrasound imaging is used routinely in its B-Mode imaging mode to report on soft tissue structures. It's also used in its Doppler modes for the measurement of blood velocity in fast-flowing targets such as the cardiovascular system, in slow-flowing applications such as quantifying blood flow and in vascular architectures within tumors.

SEM observation of a specimen cross section can provide important information for research and development as well as failure analysis. In most cases, surface observation alone cannot provide information concerning the cross sectional structure of granular materials, layered materials, fibrous materials, and powders. Preparing highly-polished cross sections of these materials is both a science and an art.

Typically, a cross section is prepared using mechanical means like conventional mechanical polishing methods or a microtome. The sample is first embedded in a holder or device, and then polished to achieve a flat cross section. In some cases, a staining procedure is used to highlight a specific component of the sample. Such methods can be lengthy procedures that require a great deal of skill, and can introduce artifacts into soft materials, deform the material around voids, or compress layers of soft and hard materials in composite samples. Mechanical polishing can miss fine details such as the presence of hairline cracks, and present a challenge to water-soluble phases.

Perfusion, washing out blood and using the open vascular channel to infuse fixative, is the standard first step of preparing animal tissues for later examination under a microscope. The rapid and homogeneous fixation resulting provides an advantage over immersion fixation that is usually used for biopsy and clinical tissue samples (Cammermeyer, 1960, Garman, 1990). A disadvantage is that brain tissue, and probably other soft tissues, prepared by perfusion has no retained extracellular space post perfusion, although living brain has about 20% extracellular space (discussed in Cragg, 1980). Perfused brain is also about 20% shrunk in whole organ volume from living brain size. The shrinkage is uneven, and distorts the relationships of structures. It is possible to avoid the organ shrinkage and distortion that follows a traditional perfusion, with little additional effort. This is a discussion of how to optimize your perfusion results, why the shrinkage occurs, and how it can be avoided.

Since the early 1990s EBSD (Electron Back Scatter Diffraction) has been developed to become a mature crystallographic analysis technique at the micro and nano-scale. It is applied in a SEM (Scanning Electron Microscope) on samples with a very smooth and clean surface. It provides quantitative orientations of individual grains, and by mapping a larger area, multiple grain analysis, texture, and grain boundaries can be examined. As the useful information for the EBSD technique comes from a very shallow depth in the material (10 to 20nm), it is a surface analysis technique showing lateral 2D distributions of crystal orientations.

One-micron plastic "thick" sections, usually stained with toluidine blue, are indispensable light microscopic adjuvants for the selection and evaluation of tissue blocks prior to electron microscopy. Plastic thick sections may also be the only practical alternative to routine histologic sections when tissue specimens are limited or too small for processing in paraffin. In all cases, however, plastic sections provide superior light microscopic resolution compared to paraffin sections, but lack the compatibility with the wide variety of special stains that make paraffin sections so versatile.

Despite the advent of the new PC based digital camera era, there is still a vast archive of film. The obvious thing to do is to scan film optically using a high resolution film scanner and to convert the image into a digital file for digital distribution, PC archiving, and image processing. The likes of NASA can easily afford the best $20k plus PMT drum scanners and pro flatbed scanners for their spaceflight photo archives. See some professional scanners at http://www.imacon.dk, http://www.aztek.com, http://graphics.kodak.com (Creo) and http://www.flatbed-scanner-review.org. You can download selections from the NASA digitised archive at http://grin.hq.nasa.gov.

High-resolution Pt/C shadowing and replication provided important insights into the size and shape of polymers beginning over 40 years ago. The first images of DNA molecules were made this way. However, in my opinion, this methodology has largely been supplanted by the use of Atomic Force Microscopy (AFM), both for direct height measurements (available in Pt/C replicas by measuring shadow lengths when the coating is deposited from one direction only) and for imaging molecular contours.

As an everyday example, consider that making a magnetic read and write head for a hard disk drive requires controlling the relative heights of several different regions to a tolerance of about 1 nm. AFM supports production by providing a rapid means of offline analysis, far faster and more precise than any replica method could be.

For years, I have used a hybrid Epon-analog resin to embed in culture dishes. I use a standard Epon formula but utilize the following components: LX-112 and DMP-30 from Ladd Research Industries DDSA and NMA from Electron Microscopy Sciences I know it seems weird, but years ago I tried all sorts things, from the “straight” formulations from each vendor to a bunch of mixtures. This one has never reacted with the plastic of the dishes. Here is how I do the actual embedding of the cell monolayers in the dishes:

1. 1) After the last 100% ethanol, remove the alcohol and cover the bottom of the well with a layer of resin mixture that is about 2 mm deep.

2. 2) Insert embedding tubes that are made by cutting the pyramidal bottoms off of BEEM capsules (just slice them with a fresh razor blade and be sure to insert them so that the manufactured end rather than the cut one is sitting against the dish).

3. 3) After inserting labels into the tubes, put them into the oven at 60° overnight.

4. 4) In the morning, fill just the embedding tubes and return everything to the oven again to finish polymerizing.

5. 5) When the resin is cured, grab the tubes with a pair of needlenosed pliers and snap them out. Sometimes a bit of the bottom of the dish comes away with the block, but often a very smooth block face results. If some of the dish comes up, it is easy to see under a dissecting microscope, and the dish portion comes away easily when trimming the block face.

I often cut away part of the block face with a jeweler's saw, either to keep it in reserve or to re-embed it in order to get cross sections, and then trim the rest into a narrow rectangle.

A challenge was presented to us to find a way to support Drosophila melanogaster so that both the anterior and posterior scutellars could be easily visualized under a dissecting microscope and photographed for purpose of measurement. The resulting solution was to utilize a standard grid box with the lid removed. Several holes were filled with Gary's Magic Mountant to just above the rim of the hole. Freshly dead fruitflies were gently lifted with forceps. Then each was placed into a filled hole and both the head and abdomen were pushed into the mountant. The wings were arranged on the plastic on either side of the hole as seen in the micrographs.

The best results were obtained if the body of the fly was evenly covered with Gary's Magic Mountant and if the photography was accomplished within a few hours of the death of the fly.

Depending on the nature of the study and what sort of information one is trying to gather through the use of FRET, FLIM has some compelling advantages in certain situations, and can provide a quantitative evaluation of the donor, acceptor and FRET pair stoichiometry. It does require access to specialized equipment and software. Different approaches to FLIM data acquisition have different strengths and weaknesses. For dynamic studies requiring high time resolution, FLIM acquisition times can fall well short of ideal.

If a yes/no answer to whether FRET is occurring is all that is required, then the polarization anisotropy of the acceptor can be used to determine FRET between fluorescent proteins (Rizzo and Piston, 2005). This is a relatively simple and robust method for confirming the presence/absence of FRET.

Precollege science education in the United States is not what it could, and should, be. Major changes are being made in the way science is taught, but delivering those changes to thousands of schools is an enormous task. Scientific societies are a major resource; they can organize and train member-volunteers to help teachers bring “real” science to the classroom. The Microscopy Society of America has become part of the effort with Project MICRO (Microscopy In Curriculum - Research Outreach). MICRO is putting MSA members, teaching materials, and microscopes in middle school classrooms nationwide. The idea began in 1993, but it has taken a lot of time and effort to implement.

MSA's early decision to collaborate with experienced science educators at the Lawrence Hall of Science of the University of California at Berkeley was a wise one; their educational materials have a well-earned national reputation for excellence.