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.

Tissue preparation for transmission electron microscopy

The basic preparation methods for routine TEM are provided in this chapter. More detailed discussions of these, plus alternative and specialized procedures, can be found elsewhere (Glauert 1972; Robards & Wilson 1993; Allen & Lawrence 1994; Glauert & Lewis 1998; Hayat 2000). A flow chart summarizing the steps required for preparing the basic range of diagnostic TEM specimens is given in Figure 22.1.

Figure 22.1 Flow chart illustrating the major steps in the preparation of specimens for diagnosis by electron microscopy.

The fundamental principle underlying TEM is that electrons pass through the section to give an image of the specimen. However, the electron beam is only capable of penetrating around 100 nm, so, to obtain a high-quality image and optimize the resolution of the instrument, it is necessary to section the tissue to a thickness of around 80 nm.

Sectioning at this level requires tissues to be embedded in extremely rigid material. Clearly the wax embedding media used in light microscopy are not suitable. In routine TEM synthetic embedding resins are used which are capable of withstanding the vacuum in the electron microscope column and the heat generated as the electrons pass through the section. Although hydrophilic media are available, in most circumstances, hydrophobic epoxy resins are preferred.

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.

Fixation

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

Formaldehyde

Commercially supplied formaldehyde solutions (i.e. formalin) normally contain some level of formic acid and considerable quantities of methanol. As such it is a poor cytological fixative and should not be used for TEM. By contrast, formaldehyde that has been freshly prepared from paraformaldehyde powder is adequate for TEM as it lacks impurities and also has the advantage of a faster penetration rate compared with glutaraldehyde. Paraformaldehyde is often recommended in electron immunocytochemistry as epitopes are less likely to be significantly altered during fixation and, if required, antigen unmasking is more effective.

Aldehyde combinations

The use of an aldehyde mixture has been proposed as a way of offsetting the disadvantages of glutaraldehyde (a slow penetration rate) and formaldehyde (less stable fixation) when applied individually (Karnovsky 1965).

Paraformaldehyde (2%) and glutaraldehyde (2.5%) fixative (buffered) (based on Glauert 1972; Karnovsky 1965)

Stock reagents

0.2 M buffer, pH 7.4 (phosphate, cacodylate)	50 ml
Paraformaldehyde	2.0 g
25% aqueous glutaraldehyde	10 ml
Distilled/deionized water to	100 ml

Method

1. Completely dissolve paraformaldehyde in buffer using heat and with continuous stirring. It may be necessary to add a few drops of 1.0 M sodium hydroxide to clarify the solution.

2. Cool the solution rapidly under running water.

3. Add aqueous glutaraldehyde. Check the pH of the mixture and adjust if necessary to pH 7.4.

4. Add distilled water to make 100 ml.

Note

Adding 0.2 ml of 1.0 M calcium chloride is thought to have a membrane stabilizing effect but this may precipitate if phosphate buffer is used.

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.

Osmium tetroxide fixative (2% aqueous)

Stock reagent

Osmium tetroxide	1.0 g
Distilled/deionized water	50 ml

Method

1. Clean, then score the glass ampoule with a diamond pencil and place in a dark-glass storage bottle.

2. Break the ampoule with a glass rod and add water. It may take 24 hours or longer for the osmium tetroxide to dissolve completely.

3. Prepared solutions may be stored for short periods at room temperature in the dark in a well-sealed bottle (double wrap the bottle in aluminum foil); for long periods store at 4°C. All osmium solutions should be stored inside a second closed container to prevent the leakage of osmium fumes. Aqueous osmium solutions that are prepared and stored in a clean container should last for ~1 year; solutions in buffer may only last a few days before they deteriorate.

4. For a 1% working solution combine 1:1 with water or buffer.

Note

Osmium is readily reduced by dust and light. Only glassware that has been acid cleaned and thoroughly rinsed in distilled water should be used. Prepared solutions should be monitored during storage and discarded if a pink color develops.

Wash buffer and staining

After primary fixation in glutaraldehyde, tissue may be treated in several ways. Material that is to be retained may be rinsed briefly (in a buffer compatible with the fixative vehicle), then stored in fresh buffer. Tissue that is for immediate processing should be washed in buffer before post-fixation in osmium tetroxide, then washed again in buffer or water to remove excess osmium. This is critical as osmium tetroxide and alcohol react to form a black precipitate. An optional step at this point is to immerse tissues after post-fixation in 2% aqueous uranyl acetate. This en bloc staining procedure adds to the contrast of the final sections and improves preservation. However, note that uranyl acetate can extract glycogen.

Dehydration

The most common embedding compounds used in TEM are epoxy resins. These are totally immiscible with water, thus requiring specimens to be dehydrated.

Dehydration is performed by passing the specimen through increasing concentrations of an organic solvent. It is necessary to use a graded series to prevent the damage that would occur with extreme changes in solvent concentration, but it is also important to keep the dehydration times as brief as possible to minimize the risk of extracting cellular constituents. The most frequently used dehydrants are acetone and ethanol. Acetone should be avoided if en bloc staining with uranyl acetate has been performed to prevent precipitation of uranium salts. Ethanol overcomes this difficulty but requires the use of propylene oxide (1,2-epoxypropane) as a transition solvent to facilitate resin infiltration. Residual dehydrant can result in soft or patchy blocks.

Commercially available ‘absolute’ ethanol normally contains a small percentage of water. This will severely restrict infiltration and polymerization of the resin and it is necessary to complete dehydration in anhydrous ethanol (which can be obtained commercially or prepared by using an appropriate molecular sieve). Propylene oxide is highly volatile, flammable and may form explosive peroxides; it should be stored at room temperature in a flammable solvents facility.

Embedding

The standard practice following dehydration and, if required, treatment with a transitional solvent, is to infiltrate the tissue sample with liquid resin. This usually requires gradual introduction of the resin, beginning with a 50 : 50 mix of transition solvent (propylene oxide) and resin followed by a 25 : 75 transition solvent resin mix, then, finally, pure resin. An hour in each of the preliminary infiltration steps is usually adequate, although it is preferable to leave samples in pure resin for 24 hours. Gentle agitation using a low-speed, angled rotator during these steps is recommended, as failure to completely infiltrate the tissue will cause major sectioning difficulties.

Once infiltrated, tissue samples are placed in an appropriate mold which is filled with resin and allowed to polymerize using heat. A paper strip bearing the tissue identification code written in pencil or laser-printed is included. Various shaped and sized molds are available. Capsules made from polyethylene are recommended as they do not react with resin, as are flat embedding molds made of silicone rubber. Polymerized blocks can be easily removed from the latter by bending the mold, which can then be reused. Polyethylene capsules can be cut away from the block using a razor or scalpel blade or the block can be extruded from the capsule using large forceps or pliers.

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).

Acrylic resins

Acrylic resins (methacrylates) derive from methacrylic acid [CH₂=C·(CH₃)COOH] and acrylic acid [CH₂=CH·COOH] and were the original synthetic media developed for use in TEM. Acrylic resins can rapidly infiltrate fixed, dehydrated tissues at room temperature. However, marked, variable shrinkage of tissue components was common due to unreliable polymerization and acrylic resins are relatively unstable in the electron beam. Currently available acrylics are now polymerized using a cross-linking process, thereby overcoming earlier disadvantages. Acrylic monomers are of low viscosity, and both hydrophilic and hydrophobic forms are obtainable. Acrylic resins react by free radical polymerization, which can be initiated using light, heat or a chemical accelerator (catalyst) at room temperature.

The main commercial acrylic resins are LR White and LR Gold and the Lowicryl series (K4M, K11M, HM20 and HM23). Each of these can be used for low-temperature dehydration and embedding to reduce the heat damage from exothermic polymerization and extraction by solvents and resin components (Acetarin et al. 1986; Newman and Hobot 1987, 1993). These characteristics make several forms of acrylic resin ideally suited to electron immunogold labeling (Stirling 1994) and enzyme cytochemical studies.

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

Cultured cells

Cell cultures may be fixed in situ, then separated from the substrate, centrifuged into a pellet and treated as a solid tissue. Alternatively, cells can be harvested into a centrifuge tube, pelleted lightly, resuspended in fixative and again pelleted by gentle centrifugation. After fixation the tube is inverted to dislodge the pellet, which is then cut into cubes for further processing. Finally cell cultures can be fixed and processed while attached to the substratum, after which inverted embedding capsules are pressed onto the cell layer. Once polymerized, blocks can be separated by force or after being cooled in liquid nitrogen (see ‘Pop-off’ technique, below).

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).

Preparing particulate specimens

Stock reagents

15% aqueous bovine serum albumen (BSA)

0.1% tannic acid (low molecular weight) in buffer, pH 7.4 (phosphate)

Method

1. Centrifuge the material in buffer in a plastic centrifuge tube to form a loose pellet.

2. Discard supernatant and resuspend the material in glutaraldehyde fixative at room temperature for a minimum of 1 hour.

3. Centrifuge the material and carefully discard the supernatant.

4. Wash the specimen by resuspending it in buffer for 10–15 minutes.

5. Centrifuge the material to form a loose pellet.

6. Discard supernatant and introduce 0.5 ml of 15% aqueous BSA. Resuspend the specimen and allow it to infiltrate for a minimum of 1 hour.

7. Centrifuge the material and discard most of the supernatant, leaving sufficient to cover the pellet to a depth of approximately 1 mm.

8. Introduce an equal volume of glutaraldehyde fixative to form a layer above the BSA. Allow material to solidify for 2–24 hours.

9. Remove the material (this is most easily achieved by cutting away the plastic centrifuge tube) and divide into small portions.

10. Wash in four changes of buffer, each for 5 minutes (for ciliary biopsies only, incubate for 15 minutes in buffered tannic acid solution, then wash in four changes of buffer, each for 5 minutes, before proceeding to step 11).

11. Postfix in 1% aqueous osmium tetroxide and process as normal.

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.

Reprocessing paraffin-embedded material

Method

1. Remove the area of interest from the block, taking care not to damage the tissue.

2. Dewax the specimen by passing through several changes of xylene. The time required depends on the size of the sample but should be at least 1 hour. A minimum of three changes is recommended.

3. Rehydrate the material in a graded ethanol series.

4. Wash in water, postfix in osmium tetroxide and process as routine specimen (see above).

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

Method

1. Remove the coverslip by soaking the slide in xylene. (This may take some time. An alternative is to firstly cool slides to −20°C for up to one hour, then carefully pry off coverslip with a blade.)

If additional fixation is required

2. Rehydrate the tissue in a graded ethanol series.

3. Wash in buffer and fix the tissue in glutaraldehyde fixative for 15–20 minutes.

4. Wash in buffer and postfix in 1% osmium tetroxide for 20–30 minutes.

5. Wash in buffer or distilled water, and then cover with 2% uranyl acetate for 15 minutes.

6. Dehydrate the tissue by passing the slide through 70%, 90%, 95%, 100% and super dry ethanol for 5 minutes in each stage.

7. Dip the slide into propylene oxide for 5 minutes. The tissue should not be allowed to dry.

8. Cover the tissue with a 2 : 1 mixture of propylene oxide and epoxy resin for 5–15 minutes. The tissue should not be allowed to dry.

9. Cover the tissue with a 1 : 2 mixture of propylene oxide and epoxy resin for 5–15 minutes. The tissue should not be allowed to dry.

10. Cover the tissue with neat epoxy resin for 5–15 minutes.

11. Drain off surplus resin mixture. Invert a freshly filled (to overflowing) embedding capsule over the section and press onto the slide.

12. Incubate the slide and capsule at 60°C for 24 hours for polymerization to occur.

13. Remove the slide and, while still warm, separate the capsule and the newly embedded tissue from the glass slide.

If additional fixation is not required

2. Dip the slide in equal parts of propylene oxide and xylene, then into propylene oxide for 5–10 minutes. The tissue should not be allowed to dry.

3. Cover the tissue with a 2 : 1 mixture of propylene oxide and epoxy resin for 5–15 minutes. The tissue should not be allowed to dry.

4. Cover the tissue with a 1 : 2 mixture of propylene oxide and epoxy resin for 5–15 minutes. The tissue should not be allowed to dry.

5. Cover the tissue with neat epoxy resin for 5–15 minutes.

6. Drain off surplus resin mixture. Invert a freshly filled (to overflowing) embedding capsule over the section and press onto the slide.

7. Incubate the slide and capsule at 60°C for 24 hours for polymerization to occur.

8. Remove the slide and, while still warm, separate the capsule and the newly embedded tissue from the glass slide.

Note

Sections (or cell cultures) are easier to prepare using the pop-off method if mounted (or grown) directly on Thermanox coverslips. (Thermanox coverslips are made of a proprietary polystyrene-like compound that is resistant to fixatives and common solvents and which separates easily from the face of resin blocks.)

Ultramicrotomy

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).

Figure 22.2 Glass knife prepared from 6.4 mm thick glass strip. Note the straight cutting edge and conchoidal fracture mark.

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).

Figure 22.3 Glass knives: left, bare knife used for trimming and semi-thin sectioning; right, knife used for ultra-thin sectioning with trough (prepared from plastic (PVC) tape) fitted.

Diamond knives

A well-maintained diamond knife is capable of cutting any type of resin block and most biological and many non-biological materials. Knives are priced according to the length of the actual cutting edge. Manufacturers supply diamond knives already mounted in a metal block (incorporating a section collection trough) designed to fit directly into the knife holder of the ultramicrotome. Diamond knives are brittle but very durable and will continue to cut for quite some time provided they are kept clean and treated carefully. The cutting edge can be cleaned by carefully running a polystyrene cleaning strip (available commercially) along (never across) the edge. A diamond knife must only be used to cut ultra-thin sections and should never be used ‘dry’ without a trough fluid.

Trough fluids

The simplest and most suitable fluid routinely used in section collecting troughs is distilled or deionized water; 10–15% solutions of ethanol or acetone can also be used (not with a diamond knife). It is important to ensure that the correct level of fluid is added. If the level is too high, the fluid will be drawn over the cutting edge and down the back of the knife, thereby preventing proper sectioning. If the level is too low, sections will accumulate on the cutting edge and will not float out.

Block trimming

Once polymerized, blocks must be cleared of excess resin to expose the tissue for sectioning. At the completion of this process the trimmed area should resemble a flat-topped pyramid with a square or trapezium-shaped face (Fig. 22.4).

Figure 22.4 Araldite blocks

(a) untrimmed, (b) trimmed of excess resin.

Trimming the block can be achieved manually or by using the ultramicrotome. At its simplest, manual trimming can be performed by mounting the block in a suitable holder under a dissecting microscope and removing the surplus resin with a single-edged razor blade. Although this method is quite speedy, considerable care is required to ensure the ultimate cutting surface is as level as possible to facilitate sectioning. Alternatively, the block is positioned in the ultramicrotome and mechanically trimmed using a glass knife.

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.

Toluidine blue stain for semi-thin sections

Stock reagents

Sodium tetraborate (borax)	1 g
Toluidine blue	1 g
Distilled water	100 ml

Dissolve the borax in the distilled water and then add the toluidine blue. After filtering, the final solution can be stored at room temperature.

Method

1. Cover sections with staining solution and heat on a hot plate at 70–80°C.

2. Allow to stain adequately (the time is not crucial – up to 60 seconds is usually sufficient), then wash thoroughly in running water. Allow section to dry.

3. Sections can be viewed dry or mounted in DPX or epoxy resin.

Note

Borax raises the pH of the final stain to around pH 11.

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.

Figure 22.5 Some examples of specimen grids.

From top left: mesh (200 size); slotted (200 size); parallel with divider (200 size); mesh (50 size); hexagonal (7 size); parallel (75 size); freeze fracture; single hole; slotted; tabbed mesh (400 size); tabbed mesh (75 size).

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.

Figure 22.6 Apparatus for application of plastic support films.

The water level is raised over the level of the wire mesh, on which grids are then placed. Approximately 0.2 ml of liquid plastic film is dropped onto the water surface over the submerged grids and the solvent allowed to evaporate. The water is then drawn off, allowing the film of plastic to settle onto the grids.

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Tags: Bancroft’s Theory and Practice of Histological Techniques

Dec 13, 2017 | Posted by admin in HISTOLOGY | Comments Off

Solution A
Disodium hydrogen phosphate (Na₂HPO₄ anhydrous)	14.2 g
Distilled water	1000 ml
Solution B
Sodium dihydrogen phosphate (NaH₂PO₄·2H₂O)	51.6 g
Distilled water	1000 ml

25% glutaraldehyde stock solution	10 ml
0.1 M phosphate buffer, pH 7.4	90 ml

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