The atomic force microscope (AFM) is well known for its outstanding spatial resolution, but it is becoming increasingly useful as the instrument for force spectroscopy. In the force spectroscopy mode, the AFM can measure tiny tension forces, in the piconewton (pN) range. Daniel Müller, Wolfgang Baurmeister, and Andreas Engel have used the AFM in both the imaging and force spectroscopy modes to pull proteins out of membranes in a controlled fashion.

Müller et al. used Deinococcus radiodurans, a bacterium best known for its high resistance to radiation (as its Genus name implies), as their test subject. They extracted a highly regular membrane from the bacterium, the hexagonally packed intermediate (HPl) layer. They mounted the HPI on mica, so that the hydrophilic outer surface of the HPI adsorbed strongly to the mica, exposing the hydrophobic inner surface to the silicon nitride AFM stylus.

This article is the second of a series transcribed from the discussion taped during the Facility Management session at M&M 2000. It is printed with permission of the Microscopy Society of America. Bulleted paragraphs indicate comments by individuals attending the session.

The expanding field of cryobiology, in particular that of the study of vitrification for long term storage of tissues for transplantation, has demonstrated that ice is damaging to smooth muscle tissue. Consequently conventional methods such as ultrarapid freezing and freeze substitution are becoming routine protocols to determine the quality of cryopreservation. This article introduces the scientific community to the PS-1000 cryofixation unit (Delaware Diamond Knives, Wilmington, DE) which provides both ultrarapid freezing and a means of validation of freeze substitution methods. When any tissue is frozen or vitrif ed for clinical use it is imperative to know the structural and functional integrity of these tissues. Ice formation within the extracellular matrix and dehydration of multi-cellular tissues, using conventional cryopreservation, is the principal reason why these methods frequently prove to be ineffective.

Over the several years that imaging energy filters have been available commercially, numerous and wide-ranging applications have demonstrated elemental mapping with a resolution approaching 1 nm. A few reports have even shown resolutions <0.4 nm. Elemental mapping by energy-filtered transmission electron microscopy (Eftem) is clearly an attractive and powerful tool, but some aspects of the techniques can be complex, with many pitfalls awaiting the unwary. This tutorial aims to cover some practical aspects of elemental mapping by Eftem. It is based largely on the author's work at the ORNL Share User Facility, where Eftem research has been performed since 1994 with a Gatan imaging filter (GIF) interfaced to a Philips CM30T operated at 300 kV with a LaB6 cathode. Most of the applications have been to metals and ceramics, emphasizing interfacial segregation and precipitation.

It has been estimated that more than 90% of all scanning electron microscope (SEM) images ever published have been obtained using secondary electrons (SE) which are defined as those electrons emitted with energies between 0 and 50 eV. The properties of these secondary electrons are therefore of considerable interest and importance. However, although secondary electrons have been intensively studied since their discovery by Starke in 1901, the majority of the work has been aimed at determining the SE yield coefficient and its variation with energy for elements and compounds. The energy spectrum of secondary electrons has received far less attention although it is evident that the form of the spectrum must have an effect on the image contrast observed in the SEM because SE detectors are energy selective devices. The few studies that have been made have mostly concentrated on spectra obtained from clean samples observed under ultra-high vacuum conditions.

We have witnessed a slow but steady decline in electron microscopy (EM) skills in the biological sciences over the last decade. This decline is illustrated by the continued closures of EM facilities throughout the world. It is, however, difficult to rationalize this decline when the need for these skills is still under a constant demand. Certainly, the introduction of new microscopes has played a part in this. High-resolution confocal light microscopes, two photon imaging and other new technologies are providing impressive amounts of new information on biological systems. Many of these approaches require simple preparation protocols providing information that can be easily interpreted and are thus easily adapted to fast-moving research.

The application of EM techniques is also deceivingly simple. For this reason the field of immunocytochemistry is undergoing an increase in popularity. Laboratories with little or no previous experience in EM are discovering the impressive and convincing results achievable using these high-resolution methods.

We do this two different ways, using polyethylene (PE) molds or aluminum weigh boats, depending on the sample. You can either heat- or UV-cure these molds – just change the type of “lid” on the embedding mold.

The PE molds are used when the coverslip can be made to fit in the mold. We use JB-4-type molds, available from your favorite microscopy supplier. The key to using these molds is that they have to be pre-treated…fill the inner cup of the mold with LR White, cover with an aluminum JB-4 chuck (try to have enough resin in the mold to come up around the base of the chuck), and polymerize in a 60° C oven for a day or two, until the resin is hard. The Al chuck can usually be removed by hand, but a flathead screwdriver used as a pry will help pop it off if you have trouble.

A very basic point that is almost always overlooked when discussing SEM “resolution,” is the difference between “resolution” and “resolving power.”

An SEM might have a resolving power of 1 nm or so based on the observation of gaps between gold particles on a featureless carbon film. Fine—if ones job is to look at gold on C.

For the actual 3-D samples most everyone looks at, EVERY micrograph has a DIFFERENT resolution, closer to 10 nm, because of secondaries generated by back scattered electrons, edge effects, …etc. If you want resolutions closer to the instrument's resolving power you have to look at thin TEM specimens without sharp edges and angles.

In our semiconductor lab we have several FESEMs advertised to have resolutions of 1 nm that can not resolve 4 and 5 nm layers in a film stack.

Make sure that gloves are worn for this procedure. All containers and gloves used should be rinsed with water and the water put in the waste mercury container for next time.

All solutions containing mercury salts are collected into a jar labelled “Used Mercury Salts”. We keep this in our grossing room. The jar is actually a clear glass 3 kg sodium phosphate container.

When the jar is about half full, to each 900 mL (estimate, don't measure) add 40 grams of sodium carbonate to raise the pH to 8.0 or higher and mix well.

Leave it for a while, then filter the solution and collect the precipitate into a plastic jar for disposal. Put the filter paper in the jar as well. Seal the jar very well. We coat the lid with several layers of paraffin wax until it is about a quarter inch thick.

Many years ago we regularly used platinum crucibles in many analytical chemistry procedures, and according to my analytical chemistry book, platinum can be cleaned by fusion with potassium or sodium bisulfite. I later adopted this procedure with good success to clean the platinum apertures for an RCA EML electron microscope.

Get asmall ceramic crucible, put a bit of the bisulfite in it, heat it until the bisulfite melts, then drop the apertures into the melt. After a few minutes allow the melt to solidify, dissolve the bisulfite with hot water, and your apertures should be as bright and shiny as new.

Diatoms make for beautiful specimens for both transmission and scanning electron microscopy. As well, they are studied by many people, and there is always a need for good diatom preparations for EM.

I find that the diatoms in diatomaceous earth are usually broken, and like to prepare my own from fresh specimens. The materials needed for this are a plankton net, some potassium dichromate, and 30% hydrogen peroxide. Once the diatoms are collected (plankton net tows from shoreline, or wherever), they can be cleaned using the chemicals.

Questions often come up concerning the best way to prepare powders for electron diffraction analysis in the TEM. This is actually fairly simple:

• 1) Take an oxide powder, grind it up with mortar and pestie and take a carbon coated grid and swipe it across the fnes. Two glass slides can also use to grind up the powder.

• 2) A number of materials can be evaporated onto a carbon coated grid or onto a cleaved NaCI sample and then float that off on water onto a grid.

• 3) A molybdenum wire can be smoked in air to provide crys tals. If the wire is left in the smoke long enough, there will probably be enough crystal to cover a grid. Heat the wire across the terminals of an evaporator or heat with a torch to generate the white smoke. Thus MoO3 sample will pro- vide a good rotation calibration sample. Magnesium can also be burned to produce MgO crystals.

Being an old timer, I have been around since before the advent of vials of glutaraldehyde. In the 1970s, the Merck index described a method for purifying glutaraldehyde using activated charcoal.

Basically, glutaraldehyde begins to polymerize when stored at room temperature. Since best fixation is achieved with the monomeric form, this purification protocol was designed to remove the more complex forms. Storage under nitrogen in a sealed vial slows the polymerization, and lower concentrations degrade more slowly. Storing at 4°C is advised.