Transmission electron microscopy

22 Transmission electron microscopy

The fundamental advantage of transmission electron microscopy (TEM) over conventional light microscopy is that the electron microscope has a resolution approximately 1000 times better than the light microscope. With this greater resolving power the transmission electron microscope is able to reveal the substructure or ultrastructure of individual cells. The physical basis of this benefit lies in the formula:


where R, the resolution, represents the capacity of the optical system to produce separate images of objects very close together, λ is the wavelength of the incident illumination, and NA is the numerical aperture of the lens.

Critically, for any given lens, resolution is directly related to the wavelength of the source radiation. For example, the limit of resolution of a bright-field microscope using glass lenses and white light is around 200 nm, whereas a fluorescence microscope operating with shorter wavelength ultraviolet light is capable of resolving objects around 100 nm apart.

By comparison, using electromagnetic lenses and a beam of electrons accelerated to a potential of 100 kV, an electron microscope is theoretically capable of resolving approximately 0.001 nm. Although flaws in lens design restrict this potential, contemporary transmission electron microscopes are capable of resolving structures of 0.2 nm or less.

Specimen handling

In order to preserve the ultrastructure of the cell it is crucial that samples are fixed as soon as possible after the biopsy is taken. The most sensitive cellular indicators of autolytic/degenerative change are the mitochondria and endoplasmic reticulum, both of which may show signs of swelling (a reflection of osmotic imbalance) only a few minutes after the cells are separated from a blood supply.

The standard approach is to immerse the specimen in fixative (preferably pre-cooled to 4°C) immediately after collection. Once in fixative, the specimen is cut into smaller samples using a scalpel or razor blade. At this point the tissue should be orientated and dissected to optimize exposure of the critical diagnostic features during sectioning and screening. Dissection must also facilitate the penetration of fixatives and processing reagents. The final tissue blocks may be in the form of thin sheets or small cubes (~1 mm3), although the risk of sampling error increases as the sample size decreases. In general, the volume of fixative should be at least 10 times the volume of the tissue. It is also vital to ensure that the tissue remains completely submerged in the fixative – small pieces may adhere to the inside of the lid of the biopsy container; these will be poorly fixed even if they have been exposed to fixative vapor. Gently agitating the vial on a mechanical rotator should help to overcome this problem and improve fixation.

The importance of using small samples cannot be overemphasized. The use of cold fixative assists in minimizing postmortem changes but fixation may be hindered as a consequence. In addition, the penetration rate of most TEM fixatives is quite slow, increasing the risk of artifact formation. It should also be noted that fixatives and processing reagents penetrate different tissues at different rates, and some tissues (such as liver) very poorly. Needle biopsies of liver may need to be cut longitudinally to ensure adequate fixation. If a delay in fixation is unavoidable, damage can be minimized by holding the tissue (for a short time only) in chilled normal saline. However, the tissue must not be frozen at any point.


The fixatives used in TEM generally comprise a fixing agent in buffer (to maintain pH) and, if necessary, with various additives to control osmolarity and ionic composition. Other factors that affect fixation include fixative concentration and temperature, and the duration of fixation. The standard protocol involves primary fixation with an aldehyde (usually glutaraldehyde) to stabilize proteins, followed by secondary fixation in osmium tetroxide to retain lipids (Hayat 1981).


Fixatives are normally buffered within the range of pH 7.2–7.6 (Robinson & Gray 1996). Ideally the osmolarity and ionic composition of the buffer should mimic that of the tissue being fixed. In general practice this is not a major requirement but, if required, 300–330 mOsm (the osmolarity equivalent to that of plasma or slightly hypertonic) is suitable for most circumstances. Non-ionic molecules such as glucose, sucrose or dextran are used to adjust tonicity as these will not influence the ionic constitution of the buffer. The addition of various salts, particularly calcium and magnesium, is thought to improve tissue preservation, possibly by stabilizing membranes (Hayat 1981). This is unlikely to have a major effect in routine diagnostic applications.

Phosphate buffers

Phosphate buffers (Gomori 1955) have the disadvantage of being good growth media for molds and other microorganisms. Additionally, most metal ions form insoluble phosphates, which restricts the use of this buffer (the phosphates of sodium, potassium and ammonium are soluble). Nevertheless, phosphate buffers are the buffer of choice as they are non-toxic and work well with most tissues.

Alternative buffers

Other buffers that have been recommended for use in TEM include cacodylate (Plumel 1948; Sabatini et al. 1963), HEPES (N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid), MOPS (3-(N-morpholino) propanesulfonic acid) and PIPES (piperazine-N,N′– bis2-ethanesulfonic acid) (Good et al. 1966; Good & Izawa 1972; Massie et al. 1972; Salema & Brandão 1973; Ferguson et al. 1980).

Aldehyde fixatives

Osmium tetroxide

The use of osmium tetroxide fixation to preserve lipids is fundamental to electron microscopy (Palade 1952; Millonig & Marinozzi 1968). While primary fixation in osmium tetroxide is effective, its extremely slow penetration rate can give rise to autolytic changes. For this reason osmium tetroxide is almost always used as a secondary fixative (termed ‘post-fixation’) after primary fixation in aldehyde. The penetration rate of osmium tetroxide is also higher in stabilized tissue, such that immersion for 60–90 minutes is usually sufficient for most specimens.

Osmium tetroxide is usually supplied in crystalline form sealed in glass ampoules. Extreme care should be exercised when preparing this material and gloves and eye protection should always be worn. It is essential to only handle osmium tetroxide in a fume-hood, as the vapor will also fix tissue.

Specimens fixed in aldehyde solutions should be washed thoroughly in buffer before post-fixation in osmium tetroxide to prevent interaction between the fixatives which can cause precipitation of reduced osmium. Osmium tetroxide can be prepared as an aqueous solution, although it can also be made in the same buffer used to prepare the primary fixative. Osmium tetroxide should be avoided if electron immunogold labeling studies are to be performed, as it has the potential to alter significantly protein structure, rendering antigenic determinants unreactive.

Epoxy resins

Epoxy resins have been the embedding medium of choice in TEM since their introduction in the mid-1950s (Glauert et al. 1956). These resins contain a characteristic chemical group in which an oxygen and two carbon atoms bond to form a three-membered ring (‘epoxide’). Cross-linking between these groups creates a three-dimensional polymer of great mechanical strength. The polymerization process generates very little shrinkage (usually less than 2%) and, once complete, is permanent. As well as their properties of uniform polymerization and low shrinkage, epoxy resins also preserve tissue ultrastructure, are stable in the electron beam, section easily and are readily available.

Epoxy resins usually comprise four ingredients: the monomeric resin, a hardener, an accelerator and a plasticizer. Although manufacturers provide advice on the appropriate proportions, the hardness and flexibility of blocks and polymerization times can be manipulated by varying the amount of the individual components. It is the proportion of each component that is important; hence resins can be prepared by volume or weight. The simplest approach is to weigh the components into a disposable paper or plastic cup; as unused resin can be polymerized and discarded in the container. Thorough mixing of the components is absolutely essential. When prepared, the resin is best delivered through a non-reactive plastic syringe or pipette.

Examples of widely used epoxy resin composites include Araldite (Glauert & Glauert 1958), Epon (Luft 1961) and Spurr’s resin (Spurr 1969). Although the original product names Araldite and Epon refer to epoxy resins developed by the CIBA Chemical Company and Shell Chemical Company, respectively, these terms are now in general use. Araldite polymers are preferred as these react with a higher degree of cross-linking and are the most stable.

Occupational exposure to epoxy resins is a common cause of allergic contact dermatitis (Kanerva et al. 1989; Jolanki et al. 1990). These agents are also probable carcinogens, primary irritants and systemically toxic (Causton 1981). Spurr’s resin in particular is toxic and should be handled with great care (Ringo et al. 1982).

Tissue processing schedules

Manual tissue processing is best performed by keeping the tissue sample in the same vial throughout, and using a fine pipette to change solutions. When processing multiple samples, take care not to cross-contaminate specimens – use separate pipettes. All vials must be clearly labeled and labels must be ‘solvent-proof’. It is advantageous to agitate tissue specimens throughout the processing cycle to enhance reagent permeation. A protocol for the routine processing of solid tissue samples is given in Table 22.1.

Table 22.1 Standard processing schedule for solid tissue cut into 1 mm3 blocks (each step is performed at room temperature unless stated otherwise)

Primary fixation 2.5% glutaraldehyde in 0.1 M phosphate buffer 2–24 hours (room temperature or 4°C)
Wash 0.1 M phosphate buffer 2 × 10 minutes on rotator
Wash Distilled water 2 × 10 minutes
En bloc staining (optional) 2% aqueous uranyl acetate 20 minutes
Dehydration 70% ethanol 10 minutes on rotator*
90% ethanol 10 minutes on rotator
95% ethanol 10 minutes on rotator
100% ethanol 15 minutes on rotator
Dry absolute ethanol 2 × 20 minutes on rotator
Transition solvent (clearing) 1,2-epoxypropane 2 × 15 minutes on rotator
Infiltration 50 : 50, clearant : resin# 1 hour
25 : 75, clearant : resin 1 hour
Resin only 1–24 hours (with vacuum to remove bubbles)
Embedding Fresh resin in embedding capsules 12–24 hours at 60–70°C

* Tissues may be stored at this stage.

# As batches may vary, resin should be prepared in accordance with manufacturer’s instructions.

Procedures for other tissue samples

Cell suspensions or particulate matter

Cell suspensions (such as fine needle biopsy aspirates, bone marrow specimens or cytology samples) or particulate materials (including fluid aspirates, tissue fragments or products and specimens for the assessment of ciliary structures) are best embedded in a protein support medium before processing. Plasma, agar or bovine serum albumen (BSA) can be used. The addition of tannic acid (Hayat 1993) during the preparation of ciliary specimens gives improved visualization of axonemal components (Sturgess & Turner 1984; Glauert & Lewis 1998). The tannic acid is thought to act as a fixative and also a mordant, facilitating the binding of heavy metal stains (Hayat 2000). Double en bloc staining with uranyl acetate and lead aspartate may also improve the visibility of dynein arms (Rippstein et al. 1987).

Material embedded in paraffin/cell smears

Occasionally it becomes necessary to examine the ultrastructure of a cell smear or specimen originally embedded in paraffin and intended for light microscopy. As the preservation quality may vary, considerable care must be exercised in the electron microscopic interpretation of such material. Nevertheless it is often possible to obtain information sufficient for diagnostic purposes.

Pop-off technique for slide-mounted sections (after Bretschneider et al.1981)


Glass knives

Knives are prepared from commercially available plate glass strips manufactured specifically for ultramicrotomy. Before use, the strips should be washed thoroughly with detergent, then rinsed in distilled water and alcohol and dried using lint-free paper. Most knifemakers will allow knives of different cutting edge angles to be produced. Higher angle knives (up to 55°) are best suited to cutting hard materials, while softer blocks respond better to shallower (35°) angle knives. Glass squares and knives should be prepared just before use to avoid contamination and stored in a dust-free, lidded box.

Knives should always be inspected before use. If the knife edge is correctly formed, when it is observed face-on it should be straight and even but with a small glass spur on the top right-hand end (Fig. 22.2).

The edge need not be horizontal, but those that are obviously convex or concave should be discarded. The knife should also display a conchoidal fracture mark that curves across and down from the top left-hand edge of the knife until it meets, and runs parallel to, the right-hand edge of the glass. Each of these characteristics is visible macroscopically. When placed in the ultramicrotome and viewed under the microscope the cutting edge will appear as a bright line against a dark background. The left third of the cutting edge should appear as a smooth line and is the zone recommended for thin sectioning. The middle third is quite frequently also adequate but can show minute imperfections, and is best reserved for trimming blocks prior to sectioning and for cutting semi-thin sections.

In ultramicrotomy, thin sections are floated out for collection as they are cut. This requires a small trough to be attached directly to the knife. Pre-formed plastic or metal troughs that can be fitted to the back of the knife are commercially available. These must be sealed with molten dental wax or nail varnish after attachment but they are expedient and simple to use. An alternative approach is to prepare a trough using self-adhesive PVC insulating tape. The lower edge of the trough so formed is then sealed with molten dental wax (Fig. 22.3).

Semi-thin sections

Semi-thin (or ‘survey’) sections allow samples to be screened for specific features and to select areas for thin sectioning. Semi-thin sections are usually cut on a glass knife. However, there are now diamond knifes specifically produced for semi-thin section microtomy.

Commonly, semi-thin sections are cut at between 0.5 and 1.0 µm from trimmed or partly trimmed blocks using the ultramicrotome and a glass knife. Sections can be cut dry (using a slow cutting speed) and picked up with forceps or directly into the flotation bath attached to the knife. Sections are transferred to a drop of water on a glass microscope slide and dried on a hot plate at 70–80°C. Semi-thin sections can be examined using phase contrast or be stained and viewed by bright-field microscopy. Various cationic dyes, including methylene blue, azure B (Richardson et al. 1960) and crystal violet, can be used for this purpose, although the most common is toluidine blue (with borax). All are applied at high alkaline pH and with heat to facilitate penetration of the resin.

Collection of sections

Ultra-thin sections are mounted onto specimen grids for viewing. Grids measure 3.05 mm in diameter and are made of conductive material, commonly copper, nickel or gold, although silver, palladium, molybdenum, aluminum, titanium, stainless steel, nylon-carbon and combination varieties are available. A large range of patterns and mesh sizes are available (Fig. 22.5), with 200 square mesh being commonly used, although slotted, parallel bar and hexagonal patterns are also standard. As electrons cannot pass through the metal grid bars, the choice of grid becomes a compromise between support for the sections (better with grids of smaller mesh size) and the relative proportion of exposed section (better with grids of larger mesh size). The latter provides a large area of section for viewing but with less stability.

Support films

The use of support films is generally unnecessary with contemporary, routine embedding media that have been properly prepared. If, however, larger viewing areas are required, it may be necessary to use support films to provide greater section stability.

Electron-transparent plastic films prepared from collodion, Formvar or Butvar are commonly used. There are many methods for applying plastic films, with one of the simpler being illustrated in Figure 22.6. The major problem with using plastic films is that the conductive properties of the grid become compromised. Re-instating the thermal and electrical properties is usually achieved by adding a 5–10 nm layer of carbon in a sputter coater or vacuum-evaporating unit.

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Dec 13, 2017 | Posted by in HISTOLOGY | Comments Off on Transmission electron microscopy

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