The atomic force microscope (AFM) has been shown to be very versatile. Fu-Ren Fan and Allen Bard have extended this versatility by imaging biologic molecules based on electrochemical reactions, referred to as scanning electrochemical microscopy (SECM). Imaging in this mode required the specimen to be covered with a thin film of water. An electrical potential of about 3 volts was maintained between the tungsten tip and an electrode near the specimen, so that electrochemical reactions that occurred within the aqueous medium resulted in a detectable current (a few picoamps).

As you can appreciate, the film of water was critical. Fan and Bard accomplished this by having the specimens on a mica chip in a environment with high relative humidity. The mica was coated with gold on one side to create the reference/counter electrode. When the SECM was operated on a dry mica chip, very small currents were measured.

Miniaturization of electronic components is now reaching the level of whole instruments, and not just parts like transistors. Thomas George recently reported on a tiny electron microscope column a “few millimeters thick and about a centimeter square”, which would work in air. The secret being an electron- transparent membrane sealing in the column's vacuum. This is exciting news for folks working in areas such as acarid chaetotaxy that would benefit from a pocket-size electron microscope. However, it's limited by a micrometer-scale resolution limit.

Help is on the way, however. In separate reports in Applied Physics Letters, Driskill-Smith et al at. published “The 'nano-triode:’ A nanoscale field-emission tube”, and Dean and Chalamala published “The environmental stability of field emission from single-walled carbon nanotubes.“

Since, as everyone knows, an electron microscope's electron gun is a triode vacuum tube, the application is obvious. Everyone old enough to remember vacuum tubes, I should say.

There was a spate of posts concerning the production and collection of ultrathin serial sections on the microscopy listserver, so I wanted to draw the microscopy community's attention to these instructions from an unfortunately obscure microscopy book. “To obtain quality serial sections for electron microscopy the following procedure must be followed for consistent results.

Microscopists have continually developed new techniques for enhancing specimen contrast since Zacharius first built the first compound light microscope around 1595. His simple two lens microscope became the basis for the evolution of light microscopes until 1828. That date is when the first polarizing light microscope was invented by Nichols. Since then, it has served as the building block for most research microscopes that are fitted for accessories made to enhance contrast or measure specific optical properties.

Julius Rheinberg devised a simple contrast enhancement method in 1896. He published his findings in the journal of the Royal Microscopical Society for the benefit of others. Today, Rheinberg illumination (RHI) has given way to phase contrast, differential interference contrast, confocal, laser microscopy, scanning probe and atomic force microscopy. The latter four are very recent technological advances.

Although scanning electron microscopes (SEM) have been used to study the polyolefin polymer morphology as it relates to the development and modification of the commercial products, it has seldom been used to study polyolefin microstructures such as spherulite and lamellae morphology. Rather, transmission electron microscopes (TEM) have been used for such morphology studies because they provide the resolution necessary for observation of polymer microstructures. Unfortunately, TEM studies require the microtomesectioning of the buik samples to produce suitable test samples after unloading. In contrast, we have shown that, if suitable sample preparation techniques are employed, the surface morphology of bulk polyolefin samples, such as the thick test pieces and commercial products, can be characterized using the SEM. In this article we introduce our chemical etching technique that is suitable for the investigation of the polyolefin microstructures.

The programmable array microscope (PAM) is a powerful tool combining the capabilities of nearly all previously described optical sectioning techniques in a single microscope. Not only can the user create optical sections of threedimensional objects, but the PAM's unique adaptive optical strategy allows a user to select the best sectioning method for a particular sample or experimental need. The key to the PAM is a spatial light modulator (SLM). This device, when placed in the image plane of a microscope, can be used to create optical sectioning, generate spatial encoding masks, and/or define regions of interest. The growing family of PAMs includes optical sectioning systems operating in fluorescence4'3'7112 and reflection7'112, as well as spectroscopic imaging systems for fluorescence emission spectroscopy. Perhaps the most common application of this unique family of adaptive optical systems is optical sectioning.

This article is the first of a series transcribed from the discussion taped during this session, it is printed with permission of the Microscopy Society of America. Bulleted paragraphs indicate comments by individuals attending the session. Additional discussion topics will be highlighted in future publications.

Sections of formalin-fixed and paraffin-embedded tissues are commonly used to detect antigens of diagnostic, therapeutic, or prognostic importance in patients with many types of cancers. Sections (- 5 μm thick) cut from paraffin-embedded tissues are mounted on glass slides usually one or two days before immunostaining. In some cases, paraffin blocks are no longer available, and so immunohistochemistry must be performed on unstained slides that have been prepared sometime ago. it is not uncommon in both clinical and research laboratories to store unstained paraffin-embedded sections on glass slides at room temperature. Also, research laboratories that serve as reference centers for multi-institutional studies store slides for future use in order to rapidly return the tissue blocks to the originating institution. Such institutions want tissue blocks returned to them as soon as possible, as this may have medicolegal implications.

Storage of sections also becomes necessary when one marker study is proceeding and future marker studies need to be followed.

Microscope immersion oil is used in light microscopy to improve imaging. The use of microscope immersion oil as part of a microscope lens system will produce a brighter and sharper image than a similar design not using immersion oil. The reason is that this oil replaces the air gaps between the condenser and the bottom of the slide and between the top of the slide or cover glass and the objective lens with a medium that has a refractive index equal to the lowest refractive index of these glass components.

How does this work? There is a cone of light extending from a centered point on the specimen to the diameter of the objective lens. The angle of this cone is the Angular Aperture (A.A.). A.A. ranges from 10° for long focus low powered objectives to 140° for short focus high power objectives.

This is not that uncommon of a question. The first rule of thumb is that if it does not look like gold, then it probably is not gold. However, there is one exception and that is when the sputtering process creates some kind of surface structure that leads to an optical effect, there may be a nongold looking gold color (more like gray). But this typically does not adversely effect the sputtering rate.

Other cases would be when either a) contamination from external sources (e.g., fnger prints) or b) build up of contaminants from the use of gold with insufficient purity. If a), then the “problem” is solved by a good solvent washing and scrubbing with something like acetone. If b), then it will not rub off with solvent and it would be something building up in the way of impurities from within the gold foil itself.

We've used gelled agarose to cover & hold monolayer cells in place when treatment leads to cell lifting. Low-gelling temperature agarose (Type VII) (Sigma A 4018 or A 0701) can be made as a 2% stock solution in “phosphate-buffered saline” (PBS) (boiled then kept at 42° C to remain liquid).

Rough trim the excess plastic from the specimen face with a razor blade, keeping all the tissue edges. Put the specimen chuck into the microtome arm, and put in a dry glass knife. Shave off enough plastic from the front (face) in very small increments (start with 1 μm slices, decrease to 0.5 μm, then 0.1 μm until you get a mirror smooth face). If you must keep all the sample, then start with a very thin shaving just to get the face smooth. You should be able to tell when you are into the tissue by looking at the thin shavings. Mark the top edge of the one that will become the top of your trapezoid-with a magic marker on the chuck.

Make up 0.2% sodium lauryl sulphate (a detergent) in distilled water. Grids are dipped in this just prior to collecting sections, then drained on filter paper before introducing under the water in the knife boat and bringing up under the sections. The sections can be manipulated along flimed slots with an eyelash before the residual film of water dries. You can play this game under the binocular microscope for some minutes while watching the water evaporate. Its a bit like watching paint dry!

Sometimes, raising the water meniscus (i.e., to convex) is useful when trying to position sections prior to collection.

Too much carry-over of detergent can cause the sections to run together and bunch up, this is a surface tension effect of the detergent.